Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications

Summary

Here we present, and contrast two protocols used to decellularize plant tissues: a detergent-based approach and a detergent-free approach. Both methods leave behind the extracellular matrix of the plant tissues used, which can then be utilized as scaffolds for tissue engineering applications.

Abstract

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor biocompatibility, and cost. Plant tissues have favorable characteristics that make them uniquely suited for use as scaffolds, such as high surface area, excellent water transport and retention, interconnected porosity, preexisting vascular networks, and a wide range of mechanical properties. Two successful methods of plant decellularization for tissue engineering applications are described here. The first method is based on detergent baths to remove cellular matter, which is similar to previously established methods used to clear mammalian tissues. The second is a detergent-free method adapted from a protocol that isolates leaf vasculature and involves the use of a heated bleach and salt bath to clear the leaves and stems. Both methods yield scaffolds with comparable mechanical properties and low cellular metabolic impact, thus allowing the user to select the protocol which better suits their intended application.

Introduction

Tissue engineering emerged in the 1980s to create living tissue substitutes, and potentially address significant organ and tissue shortages¹. One strategy has used scaffolds to stimulate and guide the body to regenerate missing tissues or organs. Although advanced manufacturing approaches such as 3-D printing have produced scaffolds with unique physical properties, the ability to manufacture scaffolds with a diverse range of achievable physical and biological properties remains a challenge^2,3. Moreover, due to a lack of a functional vascular network, these techniques have been limited in regenerating 3-dimensional tissues. The use of decellularized animal and human tissues as scaffolds has aided in circumventing this problem^4,5,6,7. However, high cost, batch-to-batch variability, and limited availability may limit widespread use of decellularized animal scaffolds⁸. There are also concerns about potential disease transmission to patients and immunologic reaction to some decellularized mammalian tissues⁹.

Cellulose, derived from plant and bacterial sources, has been extensively used to generate biomaterials for a wide range of applications in regenerative medicine. Some examples include: bone^10,11, cartilage^12,13,14 and wound healing¹⁵. Scaffolds that are comprised of cellulose have an added benefit in that they are durable and resistant to being broken down by mammalian cells. This is due to the fact that mammalian cells do not produce the enzymes necessary to break down cellulose molecules. In comparison, scaffolds produced using macromolecules from the extracellular matrix, such as collagen, are readily broken down¹⁶ and may not be well suited to long-term applications. Collagen scaffolds can be stabilized by chemical cross-linking. However, there is a trade-off due to the inherent toxicity of the cross-linkers that affect the biocompatibility of the scaffolds¹⁷. Conversely, cellulose has the potential to remain present at the site of implantation for prolonged periods of time because it is impervious to enzymatic degradation by mammalian cells^18,19,20. This can be altered by tuning the rate of degradation through hydrolysis pretreatment and co-delivery of the scaffolds with cellulases²¹. The biocompatibility of decellularized plant-derived cellulose scaffolds in vivo has also been demonstrated in a study done on mice²².

Through hundreds of millions of years of evolution, plants have refined their structure and composition to increase the efficiency of fluid transport and retention. Plant vascular vessels minimize hydraulic resistance by branching into smaller vessels, similar to the mammalian vasculature according to Murray's law²³. After decellularization, the plant's complex network of vessels and interconnected pores is maintained. Considering the vast number of distinct plant species readily available, plant-derived scaffolds have the potential to overcome design limitations currently affecting scaffolds in tissue engineering^24,25. For instance, Modulevsky et al. demonstrated that angiogenesis and cell migration occurred when decellularized apple tissue was implanted subcutaneously on the back of a mouse²². Similarly, Gershlak et al. showed that endothelial cells could be grown within the vasculature of decellularized leaves²⁴. In a separate experiment, Gershlak et al. were also able to show that cardiomyocytes could be grown on the surface of leaves and were able to contract²⁴.

Plants also include complex organization from the cellular to the macroscopic scale, which is difficult to achieve even with the most advanced manufacturing techniques developed to date. The complex hierarchical design of plant tissues makes them stronger than the sum of their constituents²⁶. Plants possess a plethora of different mechanical properties ranging from rigid and tough components such as stems, to much more flexible and pliable ones such as leaves²⁷. Leaves vary depending on species in terms of size, shape, break strength, the degree of vascularization, and can carry different degrees of hydrophilicity. Overall, these plant properties suggest that decellularized plants can serve as unique and highly functional medical devices, including as tissue engineering scaffolds.

This protocol focuses on two methods to decellularize plant tissues, such as leaves and stems, for use as scaffolds in tissue engineering. The first method is a detergent-based technique that uses a series of baths to remove DNA and cellular matter, which has been adapted from a widely used technique to decellularize mammalian and plant tissues^{6,22,25,28,29,30}. The second method is detergent-free and is adapted from a "skeletonization" protocol generally used to remove the soft tissues of leaves³¹. Prior work showed that simmering leaves in a bleach and sodium bicarbonate solution facilitated separation of the vasculature from the surrounding soft tissue³¹. This technique can be cited back to experiments carried out in the 17^th and 18^th centuries, such as the work of Albertus Seba³² and Edward Parrish³³. These experiments centered around leaving plant matter, such as leaves and fruit, submerged in water for extended periods of time (weeks to months) and allowing the softer tissues to decay away naturally. Here the "skeletonization" approach is adapted to use milder conditions, such as longer incubation times at lower temperatures, to remove cellular residues and to avoid significantly disrupting the soft tissue structure. For the experiments detailed herein, three plant types were used: Ficus hispida, Pachira aquatica and a species of Garcinia. Results of DNA quantification, mechanical tests, and impact on cellular metabolic activity from both methods are described.

Protocol

1. Decellularization of Plant Tissue Using the Detergent-based Approach

1. Use fresh or frozen F. hispida, leaf samples. Freeze unused fresh samples in a -20 °C freezer and store for future use (up to a year).
   NOTE: Use stem or leaf tissue of nearly any desired plant. Extended storage times can cause damage to the tissues.
   1. Determine the size and shape of samples to be processed on the basis of the sample’s intended use (i.e. samples cut into strips are well suited for mechanical testing applications, meanwhile 8 mm disc samples are useful in multi-well culturing applications). Cut the leaf into 8 mm discs with a sharp, clean biopsy punch while submerged under room temperature (20-25 °C) deionized H₂O.
      Note: This protocol can be used on whole leaves and stems. However, smaller samples will decellularize faster.
   2. Incubate the samples for 5-10 min at room temperature (20-25 °C) deionized H₂O on a shake plate set to a low speed setting to wash and/or thaw them. Use enough deionized H₂O to ensure all samples are thoroughly wetted.
2. Prepare a solution of 10% (w/v) sodium dodecyl sulfate (SDS) in deionized H₂O. Place the samples in a suitable container (a glass or plastic dish is ideal) and add SDS solution to completely cover the samples. Incubate samples for 5 days at room temperature (20-25 °C) on a shake plate set to a low speed to prevent damage to the samples.
   NOTE: Do not overcrowd the container as this can slow down the decellularization process and lead to uneven treatment by the SDS. During this step, samples should acquire a brown hue.
   1. After 5 days, replace the SDS solution with deionized H₂O. Incubate the samples for an additional 10-15 min on the shake plate to thoroughly rinse off any residual SDS solution.
3. Prepare a 1% (v/v) non-ionic surfactant in 10% (v/v) bleach solution. To make a 500 mL solution, mix 5 mL of the non-ionic surfactant with 50 mL of bleach then add 445 mL of deionized H₂O. Submerge samples in the freshly prepared solution.
   NOTE: The non-ionic surfactant/bleach solution does not have a long shelf life, therefore, should be used within 48 h of preparation.
4. Replace the non-ionic surfactant/bleach solution every 24 h until the samples are completely cleared (refer to Figure 1A for visual comparison). Then incubate the sample in deionized water on a shake plate for 2 min to rinse off the excess non-ionic surfactant/bleach solution. Set the shake plate to a low setting to prevent damaging the samples.
   NOTE: The time required to clear the samples used varies, depending on the type and species of the plant. The complete discoloring of the samples is indicative of full decellularization (perform DNA quantification to ensure thorough clearing).
   1. Lyophilize the samples to store them up to a year (flash freezing using liquid nitrogen is preferred, freezing samples in -80 °C is also acceptable) and store them at room temperature (20-25 °C) in low humidity.
   2. Reconstitute samples in Tris-HCl (10 mM, pH 8.5) using enough to coat samples. Rinse samples 2-3 times in serum-free media gently using a micropipette before use (i.e. use 150-300 µL per rinse, in a standard 24 well plate).
      NOTE: Tris-HCl buffer and serum-free media (i.e. DMEM, but any basic cell culture media can be substituted) are more effective in lowering the impact to cell viability of SDS treated samples than those treated with deionized H₂O alone.

2. Preparation of Samples Using the Detergent-free Decellularization Approach

NOTE: The initial steps of this procedure coincide with steps 1.1-1.1.2 (see above).

1. Prepare a 5% (v/v) bleach (NaClO) and 3% (w/v) sodium bicarbonate (NaHCO₃) solution. Warm the solution in a fume hood to 60-70 °C for F. hispida leaf samples cut into 8 mm discs while stirring on a hot plate.
   NOTE: The sodium bicarbonate can be substituted with sodium carbonate (Na₂CO₃), or sodium hydroxide (NaOH). The desired temperature varies greatly (from room temperature to 90 °C) and should be adjusted according to the properties of the samples used.
2. Once the solution reaches the desired temperature range, submerge the samples and reduce the stirring speed to avoid damaging them. After the samples are visibly cleared (refer to Figure 1B for visual comparison), remove from the bath carefully. Incubate samples once in deionized H₂O for 1-2 min to remove excess bleach solution.
   NOTE: The time required to clear the samples can vary widely. For example, cut parsley can be cleared in 10-15 min, while thicker and/or larger samples such as whole leaves or stems can take hours in high-temperature baths to clear completely.
3. Lyophilize the samples and store them at room temperature (20-25 °C) in low humidity.

Results

Both methods yielded scaffolds that were suitable for cell culture and tissue engineering applications. Figure 1 shows the general workflow of the decellularization process using an intact leaf for the detergent-based method and cut samples (8 mm diameter) for the detergent-free method. Successful decellularization of Ficus hispida tissues following both methods yielded clear and intact samples (Figure 1A and 1B<...

Discussion

Herein, two methods to decellularize plant tissues are described. The results presented here, coupled with the results of prior studies²⁵, suggest that the protocols put forth are likely applicable to a wide spectrum of plant species and can be performed on both stems and leaves. These procedures are simple and do not require specialized equipment, so plant decellularization can be carried out in most laboratories. It is noteworthy that after decellularization, the scaffolds must be functionalized...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Sodium dodecyl sulfate	Sigma Life Science	75746-1KG
Triton X-100	MP Biomedicals, LLC	807426	Non-ionic surfactant referenced in paper. Very viscous reagent, can help to cut end of pipette tip when drawing it up.
Concentrated bleach (8.25% sodium hypochlorite)	Clorox	Item #: 31009	Standard concentrated bleach.
Sodium bicarbonate	Acros Organics	217120010	Can be substituted with sodium hydroxide or sodium carbonate.
8 mm Biopunch	HealthLink	15111-80	Cuts samples that fit well in 24 well plate
Belly Dancer-Shake table	Stovall Life Sciences	BDRAA115S	Use low speeds to not damage tissues. Can use any model/brand of shake table.
Isotemp hot/stir plate	Fisher Scientific		Can use any style/brand of hot/stir plate.
Beaker	Any		Can use any size beaker as long as it will fit your samples and not overcrowd them.
Tris Hydrochloride	Fisher Scientific	BP153-500
DMEM	Corning	MT50003PC
Quant-iT Picogreen dsDNA assay	Life Technologies	P11496	Can use any dsDNA quantification mehtod on hand.