The authors would like to thank the EPSRC (EP/L016346/1), MRC, and the Doctoral Training Alliance Biosciences for Health for funding and supporting this work.

NOTE: See the Table of Materials for details related to all materials, reagents, equipment, and software used in this protocol.
1. Preparation of the fluid gel suspension bed
<ol>
	<li>Prepare 1,000 mL of dispersed agarose (0.5% w/v) by adding 5 g of agarose powder to 1,000 mL of ultrapure water (Type 1, &#62;18 m&#937;&#183;cm-1) in a 2,000 mL glass bottle.</li>
	<li>Add a 70 mm magnetic stirrer bar to the aqueous mixture and secure the bottle cap by first fully tightening and then loosening by a quarter turn.</li>
	<li>Dissolve and sterilize the mixture by placing the glass bottle into the basket of the autoclave, closing the lid, and running a cycle for 15 min at 121 &#176;C and 1 Bar. 
	NOTE: This protocol is always used for autoclaving solutions in further steps.</li>
	<li>Remove the bottle from the autoclave once the autoclave has cooled to 80 &#176;C and place it on a magnetic stirrer (unheated), with the stirring set to 800 rpm. 
	CAUTION: The bottle and the liquid remain hot.</li>
	<li>Cool the sol under ambient conditions, while maintaining constant stirring, until the temperature is lower than its Tgel (gelling point), 32 &#176;C.</li>
	<li>Remove the bottle from the stirrer and store it at 4 &#176;C. 
	&#8203;NOTE: The fluid gel can be stored until needed.</li>
</ol>
2. Preparation of bioinks
<ol>
	<li>Prepare a gellan-based bioink using a 1% (w/w) sol of low acyl gellan gum in ultrapure water (Type 1, &#62;18 m&#937;&#183;cm-1).
	<ol>
		<li>Weigh out 0.5 g of gellan powder into a weighing boat.</li>
		<li>Add 49.5 g of ultrapure water into a 100 mL glass bottle, along with a magnetic stirrer.</li>
		<li>Fold the weighing boat containing gellan power in half and add the powder to the water slowly, while constantly stirring.</li>
		<li>Dissolve and sterilize the sol using an autoclave and allow it to cool to 20 &#176;C.</li>
		<li>Store the bioink at 4 &#176;C until further use.</li>
	</ol>
	</li>
	<li>Prepare a pectin-collagen blended bioink in ultrapure water (Type 1, &#62;18 m&#937;&#183;cm-1).
	<ol>
		<li>Prepare stock 5% (w/v) low-methoxy pectin solutions by weighing out 2.5 g of pectin powder into a weighing boat.</li>
		<li>Add 50 mL of ultrapure water into a 100 mL glass bottle, along with a magnetic stirrer.</li>
		<li>Fold the weighing boat containing pectin power in half and add the powder to the water slowly, while constantly stirring.</li>
		<li>Autoclave the aqueous mixture and cool to 20 &#176;C.</li>
		<li>Prepare 1:1 and 2:1 pectin-collagen blends by either adding 3 mL of the pectin solution to 3 mL of collagen solution or by adding 4 mL of the pectin solution to 2 mL of collagen solution, respectively. Gently mix the blends using a pipette, by withdrawing and dispensing the mixture 10x. 
		NOTE: This procedure is best undertaken using cold materials on ice to prevent premature gelation of the collagen. Precooling of pectin and collagen can be achieved by storing at 4 &#176;C prior to mixing.</li>
		<li>Store at 4 &#176;C until further use.</li>
	</ol>
	</li>
	<li>Prepare an alginate-collagen blended bioink in ultrapure water (Type 1, &#62;18 m&#937;&#183;cm-1).
	<ol>
		<li>Weigh out 2 g of alginate powder into a weighing boat.</li>
		<li>Add 50 mL of ultrapure water into a 100 mL glass bottle, along with a magnetic stirrer.</li>
		<li>Fold the weighing boat containing alginate power in half and add the powder to the water slowly, while constantly stirring.</li>
		<li>Heat the dispersion to 60 &#176;C, under constant stirring, until the alginate is fully dissolved (clear, slightly brown liquid), and then cool to 20 &#176;C.</li>
		<li>Dilute the alginate solution with cell culture medium such as Dulbecco&#39;s modified eagle medium (DMEM) by adding 25 mL of the alginate solution to 25 mL of DMEM.</li>
		<li>Prepare alginate-collagen blends (1:1) by adding 3 mL of the alginate/DMEM solution to 3 mL of collagen solution. Gently mix the blends using a pipette by withdrawing and dispensing the mixture 10x and store at 4 &#176;C. 
		&#8203;NOTE: This procedure is best undertaken using cold materials on ice to prevent premature gelation of the collagen. Precooling of pectin and collagen can be achieved by storing at 4 &#176;C prior to mixing.</li>
	</ol>
	</li>
</ol>
3. Rheological characterization of bioinks
<ol>
	<li>Turn on the rheometer, insert 40 mm serrated geometries, and allow to stand for 30 min.</li>
	<li>Zero the gap height of the rheometer using the zero-gap height function.</li>
	<li>Add ~2 mL of sample on the bottom plate and lower the top geometry to create a gap height of 1 mm.</li>
	<li>Trim the sample by removing excess material expelled from between the plates. To do this, use a flat, non-abrasive edge to pull excess fluid away from the gap and soak up with tissue paper. 
	NOTE: Steps 3.2-3.4 are repeated to change the sample before each of the following steps.</li>
	<li>Undertake viscometry profiles to determine the injectability of the bioink.
	<ol>
		<li>Select viscometry test from the user options.</li>
		<li>Input the parameters for a shear rate-controlled ramp test: 0.1 to 500 s-1, with a 1 min ramp time.</li>
		<li>Repeat the viscometry ramp test on new samples under stress control, using the upper and lower stresses determined from the shear rate-controlled ramp test in step 3.5.2.</li>
	</ol>
	</li>
	<li>Undertake small deformation tests to determine the gelling characteristics of the bioink.
	<ol>
		<li>Select oscillatory testing from the user options.</li>
		<li>Input parameters into a single frequency test under constant strain: frequency 1 Hz, strain 0.5% over 1 h, while the ink gels.</li>
	</ol>
	</li>
	<li>Undertake in situ amplitude and frequency measurements on gelled samples.
	<ol>
		<li>Select oscillatory test from the user options.</li>
		<li>Select amplitude sweep and input the parameters for an amplitude sweep test that is strain-controlled: 0.01 to 500%, at a constant 1 Hz frequency.</li>
		<li>Load a new sample and select oscillatory test from the user options. Then, select frequency test, and input frequency parameters between 0.01 and 10 Hz and a strain that is within the linear viscoelastic region (LVR) of the spectra determined from the amplitude sweep data obtained in step 3.7.2 (typically a value between 50% and 80% of the LVR).</li>
	</ol>
	</li>
</ol>
4. Designing and printing 3D structures using a 3D bioprinter
<ol>
	<li>Launch CAD Software to start the generation of a CAD model.
	<ol>
		<li>Select Tools | Materials in the CAD software to define the printing parameters for the chosen bioink.</li>
		<li>Input printing parameters relevant to the printer being used; for example, for the 3D Discovery, input the estimated filament diameter (~200-500 &#181;m for most bioinks) in the thickness tab to determine the Z thickness of each layer. 
		NOTE: Delamination of the final construct is indicative of a need to increase the thickness value, while loss of resolution highlights the need to reduce thickness.</li>
		<li>Design the desired structure layer by layer using the Layer tabs in the software. Group the layers using the Group tab and assign each layer to a level on the Z plane using the Level tab.
		<ol>
			<li>For example, to generate a lattice structure (using an alginate-collagen blended bioink prepared in step 2.3), create one layer with the filaments along the x axis and a second layer with the filaments along the y axis. Assign both to a separate Level.</li>
		</ol>
		</li>
		<li>Under the Group tab, determine the build height by selecting the number of repeated units in the structure.</li>
		<li>Click the Generate tool to create a G-code for the design and view a 3D render of the structure.</li>
	</ol>
	</li>
	<li>Close BioCAD and launch the 3D Discovery human-machine interface (HMI) software to initiate the printing process.
	<ol>
		<li>Assemble the printhead according to the manufacturer&#39;s instructions. Mount the microvalve to the printhead and screw in the chosen extrusion nozzle.</li>
		<li>Click the Needle Length Measurement function to calibrate the printhead.</li>
		<li>Load a culture vessel (e.g., a 6-well plate) onto the printing platform.</li>
		<li>Aliquot the bioink into the printing cartridge and screw into the printhead above the micro-valve.</li>
		<li>Connect the assembled printhead to the pneumatic pressure system and select printhead on the HMI to engage.</li>
		<li>Click check pressure to allow tuning of the extrusion pressure.</li>
		<li>Once an appropriate pressure has been selected (~30-120 kPa depending upon the desired resolution), open the G-code generated previously and click Run to initiate the printing process.</li>
	</ol>
	</li>
</ol>
5. Preparation of skin analogues
<ol>
	<li>Culture human dermal fibroblasts (HDFs) and adipose-derived stem cells (ADSCs) in DMEM supplemented with fetal bovine serum (FBS) (10%), HEPES buffer (2.5%), and penicillin/streptomycin (1%) in T75 flasks until 90% confluency is reached. Culture human epidermal keratinocytes (HEKs) in keratinocyte growth media (KGM) until 70%-80% confluency is reached. Keep all cells under conditions of 37 &#176;C, 5% CO2, and 95% air in an incubator during culture.</li>
	<li>For the preparation of HDFs and ADSCs for dermal and adipose bioinks, wash the cells by gently pipetting 3 mL of phosphate-buffered saline (PBS) into flasks, tilt the flask to swirl the PBS over the cells, and aspirate, taking care not to disturb the attached cells.
	<ol>
		<li>To lift the cells, pipette 3 mL of 1x cell dissociation enzyme into the flask to cover the cells and place the flask in an incubator for 3 min, firmly tapping the flask against the palm of the hand to dislodge the cells. Neutralize the action of the enzyme using 6 mL of complete DMEM. 
		NOTE: Incubate for a further 2 min if the cells remain attached after tapping.</li>
		<li>For the preparation of the dermal and adipose bioinks, pipette the cell suspensions into separate 15 mL tubes and take 10 &#181;L from each for cell counting using a hemocytometer. Centrifuge the remaining cell suspensions at 300 &#215; g for 5 min to pellet the cells.</li>
		<li>Aspirate the supernatant, being careful not to disturb the pellet, add the appropriate polymer solution (prepared in step 2.2), and mix using gentle stimulation by pipetting at the following densities:
		<ol>
			<li>For the adipose layer, pipette 5 &#215; 105 ADSCs mL-1 per 1:1 collagen to pectin blend.</li>
			<li>For the papillary layer, pipette 3 &#215; 106 HDFs mL-1 per 2:1 collagen to pectin blend.</li>
			<li>For the reticular layer, pipette 1.5 &#215; 106 HDFs mL-1 per 2:1 collagen to pectin blend.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>To print, load each bioink into a separate cartridge and print the construct in the supporting fluid gel in a glass Petri dish according to the instructions in section 4.
	<ol>
		<li>Once printing is complete, inject 2 mL of 200 mM CaCl2&#8729;2H2O around the construct and 3 mL of adipogenic media (complete DMEM supplemented with 500 &#181;M isobutyl-methylxanthine [IBMX], 50 &#181;M indomethacin, and 1 &#181;M dexamethasone) into the fluid gel using a syringe and needle. Place in an incubator overnight.</li>
		<li>The following day, remove the construct from the support bath using a spatula, gently wash in PBS, and culture for 14 days in adipogenic medium in a 6-well plate.</li>
		<li>After 14 days, remove enough medium to create an air-liquid interface at the surface of the construct and seed 2 &#215; 106 keratinocytes atop the construct to create an epidermal layer.</li>
		<li>Culture further for 1 week prior to analysis.</li>
	</ol>
	</li>
</ol>
6. Preparation of carotid artery model
<ol>
	<li>Load the gellan gum bioink solution (as prepared in step 2.1) into a printer cartridge.</li>
	<li>Print the carotid artery model within a Petri dish containing the fluid gel support material according to the printing instructions in section 4.</li>
	<li>Once printing is complete, inject 2 mL of 200 mM CaCl2&#8729;2H2O around the construct using a syringe and needle.</li>
	<li>After a minimum of 3 h, remove the construct from the support bath using a spatula and gently wash in PBS.</li>
</ol>

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N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+next-generation+bioinks:+Tuning+material+properties+pre-+and+post-printing+to+optimize+cell+viability.">Toward next-generation bioinks: Tuning material properties pre- and post-printing to optimize cell viability.</a> MRS Bulletin. 42 (8), 563-570 (2017).</li><li>Mosadegh, B., Xiong, G., Dunham, S., Min, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+progress+in+3D+printing+for+cardiovascular+tissue+engineering.">Current progress in 3D printing for cardiovascular tissue engineering.</a> Biomedical Materials. 10 (3), 034002 (2015).</li><li>Zhou, K., Sun, Y., Yang, J., Mao, H., Gu, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrogels+for+3D+embedded+bioprinting:+a+focused+review+on+bioinks+and+support+baths.">Hydrogels for 3D embedded bioprinting: a focused review on bioinks and support baths.</a> Journal of Materials Chemistry B. 10 (12), 1897-1907 (2022).</li><li>Lee, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+bioprinting+of+collagen+to+rebuild+components+of+the+human+heart.">3D bioprinting of collagen to rebuild components of the human heart.</a> Science. 365 (6452), 482-487 (2019).</li><li>Kyle, S., Jessop, Z. M., Al-Sabah, A., Whitaker, I. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Printability'+of+candidate+biomaterials+for+extrusion+based+3D+printing:+state-of-the-art.">Printability' of candidate biomaterials for extrusion based 3D printing: state-of-the-art.</a> Advanced Healthcare Materials. 6 (16), 1700264 (2017).</li><li>Hinton, T. J., Lee, A., Feinberg, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+bioprinting+from+the+micrometer+to+millimeter+length+scales:+Size+does+matter.">3D bioprinting from the micrometer to millimeter length scales: Size does matter.</a> Current Opinion in Biomedical Engineering. 1, 31-37 (2017).</li><li>Seoane-Via&#241;o, I., Trenfield, S. J., Basit, A. W., Goyanes, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translating+3D+printed+pharmaceuticals:+From+hype+to+real-world+clinical+applications.">Translating 3D printed pharmaceuticals: From hype to real-world clinical applications.</a> Advanced Drug Delivery Reviews. 174, 553-575 (2021).</li><li>Jovic, T. H., Combellack, E. J., Jessop, Z. M., Whitaker, I. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+bioprinting+and+the+future+of+surgery.">3D bioprinting and the future of surgery.</a> Frontiers in Surgery. 7, 609836 (2020).</li></ol>

The authors have no conflicts of interest to declare.

Consideration of the selection of materials used for the supporting bed 
During the development stage, there were various characteristics required of the supporting bed. These characteristics included: i) maintaining sufficient structure to suspend the extruded material; ii) shear thinning capacity to allow the printhead to move freely through the supporting material; iii) rapid restructuring (self-healing properties), forming support around the deposited bioink; iv) thermally stable at both room temperature and physiological temperatures; v) neutral (i.e., uncharged) material that is relatively bioinert, across a range of pH and electrolytes (ionic species and concentrations), preventing interactions with cells and charged bioinks; vi) non-toxic; and, vii) preferably from a non-animal source.
Although there are many biopolymer materials that maintain several of these inherent characteristics, with a capacity to perform suspended 3D additive manufacture without conforming to all of these characteristics11,26,27, the intention here was to produce a supporting bed that would overcome certain practical issues associated with other supporting materials. Due to the chemical properties of agarose and, in particular, when formulated as a particulate fluid gel, all of these characteristics could be obtained. This enabled a supporting bed that could be used across a wide variety of bioinks23,24,25,28. Indeed, the bioinert nature of the material provided the potential to maintain the printed structure in situ throughout culture, allowing sufficient timescales for many different bioinks to fully develop, without changes to the biology. Furthermore, the particulate, thinning nature allowed ease of removal from the final printed construct, while the non-toxic, non-animal origin allows the potential for rapid translation toward the clinic, overcoming barriers relating to ethical and regulatory requirements.
Considerations in the selection of materials for the bioinks 
In direct-extrusion bioprinting, bioinks are deposited onto a 2D print bed. It is beneficial that monomer bioink solutions have shear-thinning behavior; however, to produce high-fidelity constructs with physiologically relevant dimensions, they must have low thixotropy and recover to sufficiently high viscosities so that they form solid filaments upon deposition29,30,31. With increased viscosity, the pressure required for extrusion is much higher, often negatively influencing the encapsulated cell viability31,32. Suspension bioprinting removes this limitation, as the extruded material is supported by the suspension bath throughout crosslinking. This development hugely increases the range of bioink formulations that can be used. For example, recent work has shown the use of low-concentration collagen solutions being printed into highly complex geometries analogous to the internal structure of the heart33,34,35. In the applications mentioned in this method, embedded printing allowed biomaterial inks to be chosen to best replicate the physiological environment for which they were intended, instead of for their ability to be printed.
Limitations in structure size 
Throughout the biofabrication literature, it has been demonstrated that different kinds of bioprinters, driven by alternative printhead technologies, may be incorporated into embedded manufacturing techniques. The technology demonstrated here is no different, with examples that include a pneumatic micro-extrusion-based bioprinter (INKREDIBLE), as demonstrated by Senior et al., and extrusion-based bioprinters with controllable microvalves (3D Discovery)23. Although this makes the technology accessible to a range of users who may already own a bioprinter, limitations on the attainable size of the structure are ultimately dependent on the bioprinter specifications in question. Initially, the main restriction upon the generation of large structures is defined by the size of the print bed, the limits of X, Y, and Z trajectories, and also the size of the vessel in which the supporting fluid gel is contained.
Limitations in resolution 
When fabricating intricate, micrometer-sized structures, the resulting resolution is highly dependent on the precision of the printer (control over step size, degree of extrusion), the internal diameter of the print nozzle, and a range of adjustable software parameters, including print speed, print pressure, and flow velocity36. In addition, control over the droplet size appears to be critical to facilitating the generation of high-resolution structures, with the best results observed in extrusion printers with a controllable microvalve. Ultimately, when all parameters are optimized, print resolutions can be achieved to match, or even be less than, the inner diameter of extrusion nozzles, with the deposited filament on the order of the micrometer scale37. This is, however, reliant on the optimization of all the printing parameters previously mentioned, and resolution can be considerably limited by the printing mechanism and precision. Pneumatic extrusion, for example, does not appear to allow for the same printing resolution as extrusion with a controllable microvalve. There is, therefore, a potential cost implication to achieve the maximum printing resolution, as such systems incur a significantly increased expense to the user.
Future outlook and potential 
At the moment, there is much interest around the use of suspended manufacturing processes to allow for the production of complex soft structures containing embedded cells, and there will undoubtedly be significant advances in the coming years. Continuous advancement in improving print resolution is given, although it remains to be seen how necessary this will be, given that the majority of biological systems are able to rearrange themselves on a molecular level. While the focus of interest in the media is around the use of 3D printed tissues to directly replace human tissues following injury or disease, any robust medical procedures enabled by these processes are some years away38,39. It is more likely that the impact of these complex culture systems will be in the screening of drugs or even used as tools, to enhance our understanding of biological processes38. In particular, developmental biology could greatly benefit here, where precise control over the special deposition of molecules will allow researchers to explore the role of multifactorial systems on tissue development processes.

Hydrogels are perfect materials to use as supports for cell growth1. Depending on the material that is used, they gel via gentle mechanisms that do not compromise cell viability2,3. The high water content (typically &#62;90%) means that nutrients and oxygen can readily diffuse into the material and waste products of cell metabolism diffuse out4. As such, cell viability has been shown to be preserved for periods in excess of 1 year5, and there are now examples of hydrogels being used to store or &#34;pause&#34; cells for future therapeutic use6. They have been widely used in tissue engineering for the production of tissue-like structures, but their use tends to be limited by the difficulty in controlling both the structure and composition of the material. Historically, hydrogel strength is comparatively low (in relation to many hard tissues), due to the high water content and low volumes occupied by the polymer matrix forming the structure. Furthermore, many routes to gelation (thermal, ionotropic, fibrillogenesis) offer reasonably slow kinetics, meaning their mechanical properties tend to develop steadily with time. With the exception of interpenetrating networks, low mechanical rigidity and slow curing times often result in bioinks that are unable to free-stand on deposition, tending to &#34;slump&#34; and lose definition when initially extruded.
In an attempt to overcome this key issue, embedded printing techniques have been developed that provide support during printing, while the mechanical properties of the construct are developing7,8. Once the microstructure of the gel has fully developed and the mechanical properties reach an optimum, the support matrix can be removed, typically through gentle washing or melting of the support phase. Initial work on this approach utilized a viscous pluronic dispersion in which the secondary phase was distributed9. More recently, Bhattacharjee et al. used gels in the form of granulated carbopol to demonstrate that arrays of cells could be suspended in the supporting gel10. Subsequently, Hinton et al. reported on the extrusion of gel-based materials containing cells into a support bed that consisted of a microgel suspension formed from granular gelatin11. After extrusion of the cell-containing hydrogel and its subsequent curing, the gelatin was removed by gentle heating of the support bath, enabling melting of the gelatin. Unfortunately, this process still presents several limitations. For example, the chemical structure of gelatin in comparison to collagen (consequently to it being a hydrolyzed form of collagen) is such that many chemical moieties across its backbone can interact with biological entities; thus, a residual support matrix could interfere with downstream biological processes. Moreover, animal-derived products pose restrictive use when looking toward the translatability of a technology. This creates challenges if the manufactured part is intended to be used clinically, or even if it is to be used to answer fundamental biological questions, where this surface contamination can cause a significant issue.
We have subsequently created a refined process that allows for the suspended manufacture of hydrogels, within a supportive matrix, that has no charge under physiological conditions and is formed from non-animal materials. Although the process can be used with a range of biopolymeric supports, agarose provides a material that is inert to biological interactions as it is sugar-based and neutrally charged at physiological pH12,13. Rather than fragmenting an already existing gel, the support material is formed through the application of shear during gelation14,15,16. This produces a matrix of particles that exhibit dendron-like features at their surface and are dispersed in a secondary matrix of ungelled polymer17,18. The result is a material with interesting material properties19,20,21,22 that can shear-thin in a similar way to previously reported granular gels, but tends to recover viscosity more rapidly when shear is removed23. Once the cell-bearing material extruded into the supporting matrix has fully matured, the support matrix can be removed through gentle agitation before being placed in culture. It has been shown that it is possible to use this process to produce materials with complex structures and recapitulate the biological structures of both skin and the osteochondral region23,24,25. This methods paper describes in detail how to manufacture the supporting material and highlights appropriate bioinks used within a variety of complex structures.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3D Discovery Bioprinter</td>
			<td>RegenHU</td>
			<td>BIOFACTORY</td>
			<td>Microvalve extrusion bioprinter</td>
		</tr>
		<tr>
			<td>Agarose&#160;</td>
			<td>Merck</td>
			<td>9012-36-6</td>
			<td>Material used to create fluid gel support baths&#160;</td>
		</tr>
		<tr>
			<td>Benchtop autoclave</td>
			<td>Prestige Medical</td>
			<td>B8L75814</td>
			<td>Classic</td>
		</tr>
		<tr>
			<td>Brilliant Blue G</td>
			<td>Merck</td>
			<td>6104-58-1</td>
			<td>Dye used to stain structures&#160;</td>
		</tr>
		<tr>
			<td>Calcium Chloride Dihydrate</td>
			<td>Merck&#160;</td>
			<td>10035-04-8</td>
			<td>Used to reticulate printed structures&#160;</td>
		</tr>
		<tr>
			<td>Dexamethasone</td>
			<td>Merck&#160;</td>
			<td>D4902-25MG</td>
			<td>Used in adipogenic media</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Merck</td>
			<td>D6429</td>
			<td>Cell culture media</td>
		</tr>
		<tr>
			<td>Duran&#160; bottle&#160;</td>
			<td>Merck</td>
			<td>Z305219</td>
			<td>glass bottle&#160;</td>
		</tr>
		<tr>
			<td>EVOS XL Core Imaging System</td>
			<td>EVOS&#8482;</td>
			<td>AMEX1000</td>
			<td>Brightfield microscope with phase contrast</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Fisher Scientific</td>
			<td>10500-064</td>
			<td>Cell culture media supplement</td>
		</tr>
		<tr>
			<td>Gellan gum</td>
			<td>Special Ingredients</td>
			<td>5060341112638</td>
			<td>Low Acyl Gellan gum used to make the bioink for the corotid artery model</td>
		</tr>
		<tr>
			<td>HEPES buffer</td>
			<td>Merck</td>
			<td>H9897-10PAK</td>
			<td>Buffer for cell culture media</td>
		</tr>
		<tr>
			<td>Indomethacin</td>
			<td>Merck</td>
			<td>I7378</td>
			<td>Used in adipogenic media</td>
		</tr>
		<tr>
			<td>Isobutyl-methylxanthine (IBMX)</td>
			<td>Merck</td>
			<td>I7018-100MG</td>
			<td>Used in adipogenic media</td>
		</tr>
		<tr>
			<td>Keratinocyte growth medium</td>
			<td>Lonza</td>
			<td>00192060</td>
			<td>Used as media to culture keratinocytes</td>
		</tr>
		<tr>
			<td>Low Methoxy Pectin</td>
			<td>CP Kelco</td>
			<td>LM-5CS</td>
			<td>Pectin used to make pectin/collagen blends</td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin</td>
			<td>Merck</td>
			<td>P4333-100ML</td>
			<td>Used to inhibit bacterial growth</td>
		</tr>
		<tr>
			<td>PureCol EZ Gel solution</td>
			<td>Merck</td>
			<td>5074</td>
			<td>Collagen solution used to make alginate/collagen blends</td>
		</tr>
		<tr>
			<td>Sodium Alginate</td>
			<td>Merck</td>
			<td>9005-38-3</td>
			<td>Alginate powder used to make alginate/collagen blends&#160;</td>
		</tr>
		<tr>
			<td>TrypLE select&#160;</td>
			<td>Fisher Scientific</td>
			<td>12563011</td>
			<td>cell dissociation enzyme</td>
		</tr>
		<tr>
			<td>T75 Flasks</td>
			<td>StarLab</td>
			<td>CC7682-4175</td>
			<td>Used for culturing cells</td>
		</tr>
	</tbody>
</table>

agarose fluid gels formed by shear processing during gelation for suspended 3d bioprinting

The use of granular matrices to support parts during the bioprinting process was first reported by Bhattacharjee et al. in 2015, and since then, several approaches have been developed for the preparation and use of supporting gel beds in 3D bioprinting. This paper describes a process to manufacture microgel suspensions using agarose (known as fluid gels), wherein particle formation is governed by the application of shear during gelation. Such processing produces carefully defined microstructures, with subsequent material properties that impart distinct advantages as embedding print media, both chemically and mechanically. These include behaving as viscoelastic solid-like materials at zero shear, limiting long-range diffusion, and demonstrating the characteristic shear-thinning behavior of flocculated systems. 
On the removal of shear stress, however, fluid gels have the capacity to rapidly recover their elastic properties. This lack of hysteresis is directly linked to the defined microstructures previously alluded to; because of the processing, reactive, non-gelled polymer chains at the particle interface facilitate interparticle interactions-similar to a Velcro effect. This rapid recovery of elastic properties enables bioprinting high-resolution parts from low-viscosity biomaterials, as rapid reformation of the support bed traps the bioink in situ, maintaining its shape. Furthermore, an advantage of agarose fluid gels is the asymmetric gelling/melting transitions (gelation temperature of ~30 &#176;C and melting temperature of ~90 &#176;C). This thermal hysteresis of agarose makes it possible to print and culture the bioprinted part in situ without the supporting fluid gel melting. This protocol shows how to manufacture agarose fluid gels and demonstrates their use to support the production of a range of complex hydrogel parts within suspended-layer additive manufacture (SLAM).

Alginate and type I collagen bioink 
Print resolution (recorded as a function of filament diameter) was observed to be directly tunable via changes in extrusion pressure (Figure 1A-C). Extrusion pressure and print resolution were directly related to the smallest filament diameters generated via printing at an extrusion pressure of 30 kPa. Interestingly, at an extrusion pressure of 30 kPa, filaments that matched the inner diameter of the extrusion nozzle could be generated (mean filament diameter: 323 &#181;m &#177; 50 &#181;m; nozzle diameter: 300 &#181;m), suggesting that a &#34;maximum resolution&#34; could be achieved. Moreover, the printing parameters for this resolution could be successfully applied to the generation of an alginate/collagen vascular tube that can be extracted and perfused (Figure 1D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64458/64458fig01.jpg" /> 
Figure 1: Generation of high-resolution prints using SLAM. Tailoring print resolution of alginate/collagen lattices as a function of filament diameter via extrusion at (A) 30 kPa, (B) 60 kPa, and (C) 120 kPa. (D) Generation of an alginate/collagen vascular tube. Scale bars = 400 &#181;m (A-C), 10 mm (D). Abbreviation: SLAM = suspended-layer additive manufacture. <a href="https://www.jove.com/files/ftp_upload/64458/64458fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Skin analogues 
SLAM was also used to create a skin-like structure (Figure 2A) using a bioink formed from a blend of collagen I and pectin. To achieve a gradient of mechanical properties similar to those found in the skin, different proportions of pectin and collagen were used in the dermal (5% w/w pectin mixed at 2:1 with a 5 mg&#8729;mL&#8722;1 collagen stock) and hypodermal (5% w/w pectin mixed at 1:1 with a 5 mg&#8729;mL&#8722;1 collagen stock) layers. The resulting structure (Figure 2Bi-Bii) was well integrated following immersion in DMEM, with no sign of delamination. Importantly, there was a high level of cell viability throughout the structure (Figure 2Biii) following a period of 14 days of culture. Interestingly, over the period of culture, the materials stiffened24, indicating a remodeling of the material.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64458/64458fig02.jpg" /> 
Figure 2: Production of a skin-like structure. (A) A schematic showing how the SLAM process was used to produce a layered structure embedded with human dermal fibroblasts. The suspending bed here was manufactured from particles formed from agarose, while the hypodermal and dermal layers were formed of varying proportions of pectin and collagen I. (B) The layered structure was intended to represent the trilayered structure of skin (i). With success in replicating this structure (ii), high levels of cell viability were noted throughout the samples, as shown by calcein-AM staining (iii).&#160;Scale bar = 5 mm (Bii). Abbreviation: SLAM = suspended-layer additive manufacture. <a href="https://www.jove.com/files/ftp_upload/64458/64458fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Carotid artery 
To push the boundaries of the method, a selection of more complex prints was produced. In one such example, a bifurcated carotid artery was printed using 1% gellan bioink (Figure 3A), which was then crosslinked by extrusion of 200 mM CaCl2 within the fluid gel bed (Figure 3B) and simply lifted from the support following gelation (Figure 3C). Despite precursor printing solutions exhibiting low viscosity, the supporting bed was successful in enabling production of the complex geometry. The artery retained its structure during deposition, crosslinking, and extraction (Figure 3), without the need to modify print codes to incorporate additional scaffolding.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64458/64458fig03.jpg" /> 
Figure 3: Fabrication process of gellan carotid artery using SLAM. (A) Extrusion of gellan within the fluid gel bed during printing, (B) completed carotid artery print within the fluid gel during crosslinking, and (C) final carotid artery model following retrieval from fluid gel support. Scale bars = 10 mm. <a href="https://www.jove.com/files/ftp_upload/64458/64458fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Agarose Fluid Gels Formed by Shear Processing During Gelation for Suspended 3D Bioprinting at JoVE.com

Agarose Fluid Gels Formed by Shear Processing During Gelation for Suspended 3D Bioprinting

Shear processing during hydrogel formation results in the production of microgel suspensions that shear-thin but rapidly restructure following the removal of shear forces. Such materials have been used as a supporting matrix for bioprinting complex, cell-laden structures. Here, methods used to manufacture the supporting bed and compatible bioinks are described.

The use of granular matrices to support parts during the bioprinting process was first reported by Bhattacharjee et al. in 2015, and since then, several ...

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2.2K Views.  University of Huddersfield. Shear processing during hydrogel formation results in the production of microgel suspensions that shear-thin but rapidly restructure following the removal of shear forces. Such materials have been used as a supporting matrix for bioprinting complex, cell-laden structures. Here, methods used to manufacture the supporting bed and compatible bioinks are described. 

Video: Agarose Fluid Gels Formed by Shear Processing During Gelation for Suspended 3D Bioprinting

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Creation of Cardiac Tissue Exhibiting Mechanical Integration of Spheroids Using 3D Bioprinting

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and fibroblasts are first isolated, counted and mixed at desired cell ratios. They are co-cultured in individual wells in ultra-low attachment 96-well plates. Within 3 days, beating spheroids form. These spheroids are then picked up by a nozzle using vacuum suction and assembled on a needle array using a 3D bioprinter. The spheroids are then allowed to fuse on the needle array. Three days after 3D bioprinting, the spheroids are removed as an intact patch, which is already spontaneously beating. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and ...

Bioprinting of Cartilage and Skin Tissue Analogs Utilizing a Novel Passive Mixing Unit Technique for Bioink Precellularization

Bioprinting is a powerful technique for the rapid and reproducible fabrication of constructs for tissue engineering applications. In this study, both cartilage and skin analogs were fabricated after bioink pre-cellularization utilizing a novel passive mixing unit technique. This technique was developed with the aim to simplify the steps involved in the mixing of a cell suspension into a highly viscous bioink. The resolution of filaments deposited through bioprinting necessitates the assurance of uniformity in cell distribution prior to printing to avoid the deposition of regions without cells or retention of large cell clumps that can clog the needle. We demonstrate the ability to rapidly blend a cell suspension with a bioink prior to bioprinting of both cartilage and skin analogs. Both tissue analogs could be cultured for up to 4 weeks. Histological analysis demonstrated both cell viability and deposition of tissue specific extracellular matrix (ECM) markers such as glycosaminoglycans (GAGs) and collagen I respectively.

Cartilage and skin analogs were bioprinted using a nanocellulose-alginate based bioink. The bioinks were cellularized prior to printing via a single step passive mixing unit. The constructs were demonstrated to be uniformly cellularized, have high viability, and exhibit favorable markers of differentiation.

Bioprinting is a powerful technique for the rapid and reproducible fabrication of constructs for tissue engineering applications. In this study, both ...

Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed Molds

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease modeling. Self-assembled tissues offer advantages over scaffold-based tissue engineering, such as enhanced matrix deposition, strength, and function. However, there are few available methods for fabricating 3D tissues without seeding cells on or within a supporting scaffold. Previously, we developed a system for fabricating self-assembled tissue rings by seeding cells into non-adhesive agarose wells. A polydimethylsiloxane (PDMS) negative was first cast in a machined polycarbonate mold, and then agarose was gelled in the PDMS negative to create ring-shaped cell seeding wells. However, the versatility of this approach was limited by the resolution of the tools available for machining the polycarbonate mold. Here, we demonstrate that 3D-printed plastic can be used as an alternative to machined polycarbonate for fabricating PDMS negatives. The 3D-printed mold and revised mold design is simpler to use, inexpensive to produce, and requires significantly less agarose and PDMS per cell seeding well. We have demonstrated that the resulting agarose wells can be used to create self-assembled tissue rings with customized diameters from a variety of different cell types. Rings can then be used for mechanical, functional, and histological analysis, or for fabricating larger and more complex tubular tissues.

This protocol describes a platform for fabricating self-assembled tissue rings in variable sizes using a customized 3D-printed plastic mold. PDMS negatives are cured in the 3D-printed mold; then agarose is cast in the cured PDMS negatives. Cells are seeded into the resulting agarose wells where they aggregate into tissue rings.

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease ...

Protocols of 3D Bioprinting of Gelatin Methacryloyl Hydrogel Based Bioinks

Gelatin methacryloyl (GelMA) has become a popular biomaterial in the field of bioprinting. The derivation of this material is gelatin, which is hydrolyzed from mammal collagen. Thus, the arginine-glycine-aspartic acid (RGD) sequences and target motifs of matrix metalloproteinase (MMP) remain on the molecular chains, which help achieve cell attachment and degradation. Furthermore, formation properties of GelMA are versatile. The methacrylamide groups allow a material to become rapidly crosslinked under light irradiation in the presence of a photoinitiator. Therefore, it makes great sense to establish suitable methods for synthesizing three-dimensional (3D) structures with this promising material. However, its low viscosity restricts GelMA's printability. Presented here are methods to carry out 3D bioprinting of GelMA hydrogels, namely the fabrication of GelMA microspheres, GelMA fibers, GelMA complex structures, and GelMA-based microfluidic chips. The resulting structures and biocompatibility of the materials as well as the printing methods are discussed. It is believed that this protocol may serve as a bridge between previously applied biomaterials and GelMA as well as contribute to the establishment of GelMA-based 3D architectures for biomedical applications.

Presented here is a method for the 3D bioprinting of gelatin methacryloyl.

Gelatin methacryloyl (GelMA) has become a popular biomaterial in the field of bioprinting. The derivation of this material is gelatin, which is hydrolyzed ...

Direct Bioprinting of 3D Multicellular Breast Spheroids onto Endothelial Networks

Bioprinting is emerging as a promising tool to fabricate 3D human cancer models that better recapitulate critical hallmarks of in vivo tissue architecture. In current layer-by-layer extrusion bioprinting, individual cells are extruded in a bioink together with complex spatial and temporal cues to promote hierarchical tissue self-assembly. However, this biofabrication technique relies on complex interactions among cells, bioinks and biochemical and biophysical cues. Thus, self-assembly may take days or even weeks, may require specific bioinks, and may not always occur when there is more than one cell type involved. We therefore developed a technique to directly bioprint pre-formed 3D breast epithelial spheroids in a variety of bioinks. Bioprinted pre-formed 3D breast epithelial spheroids sustained their viability and polarized architecture after printing. We additionally printed the 3D spheroids onto vascular endothelial cell networks to create a co-culture model. Thus, the novel bioprinting technique rapidly creates a more physiologically relevant 3D human breast model at lower cost and with higher flexibility than traditional bioprinting techniques. This versatile bioprinting technique can be extrapolated to create 3D models of other tissues in additional bioinks.

The goal of this protocol is to directly bioprint breast epithelial cells as multicellular spheroids onto pre-formed endothelial networks to rapidly create 3D breast-endothelial co-culture models which can be used for drug screening studies.

Bioprinting is emerging as a promising tool to fabricate 3D human cancer models that better recapitulate critical hallmarks of in vivo tissue ...

3D Bioprinting of Murine Cortical Astrocytes for Engineering Neural-Like Tissue

Astrocytes are glial cells with an essential role in the central nervous system (CNS), including neuronal support and functionality. These cells also respond to neural injuries and act to protect the tissue from degenerative events. In vitro studies of astrocytes&#39; functionality are important to elucidate the mechanisms involved in such events and contribute to developing therapies to treat neurological disorders. This protocol describes a method to biofabricate a neural-like tissue structure rich in astrocytes by 3D bioprinting astrocytes-laden bioink. An extrusion-based 3D bioprinter was used in this work, and astrocytes were extracted from C57Bl/6 mice pups&#39; brain cortices. The bioink was prepared by mixing cortical astrocytes from up to passage 3 to a biomaterial solution composed of gelatin, gelatin-methacryloyl (GelMA), and fibrinogen, supplemented with laminin, which presented optimal bioprinting conditions. The 3D bioprinting conditions minimized cell stress, contributing to the high viability of the astrocytes during the process, in which 74.08% &#177; 1.33% of cells were viable right after bioprinting. After 1 week of incubation, the viability of astrocytes significantly increased to 83.54% &#177; 3.00%, indicating that the 3D construct represents a suitable microenvironment for cell growth. The biomaterial composition allowed cell attachment and stimulated astrocytic behavior, with cells expressing the specific astrocytes marker glial fibrillary acidic protein (GFAP) and possessing typical astrocytic morphology. This&#160;reproducible protocol provides a valuable method to biofabricate 3D neural-like tissue rich in astrocytes that resembles cells&#39; native microenvironment, useful to researchers that aim to understand astrocytes&#39; functionality and their relation to the mechanisms involved in neurological diseases.

Here we report a method of 3D bioprinting murine cortical astrocytes for biofabricating neural-like tissues to study the functionality of astrocytes in the central nervous system and the mechanisms involving glial cells in neurological diseases and treatments.

Astrocytes are glial cells with an essential role in the central nervous system (CNS), including neuronal support and functionality. These cells also ...

Large-Scale, Automated Production of Adipose-Derived Stem Cell Spheroids for 3D Bioprinting

Adipose-derived stromal/stem cells (ASCs) are a subpopulation of cells found in the stromal vascular fraction of human subcutaneous adipose tissue recognized as a classical source of mesenchymal stromal/stem cells. Many studies have been published with ASCs for scaffold-based tissue engineering approaches, which mainly explored the behavior of these cells after their seeding on bioactive scaffolds. However, scaffold-free approaches are emerging to engineer tissues in vitro and in vivo, mainly by using spheroids, to overcome the limitations of scaffold-based approaches.
Spheroids are 3D microtissues formed by the self-assembly process. They can better mimic the architecture and microenvironment of native tissues, mainly due to the magnification of cell-to-cell and cell-to-extracellular matrix interactions. Recently, spheroids are mainly being explored as disease models, drug screening studies, and building blocks for 3D bioprinting. However, for 3D bioprinting approaches, numerous spheroids, homogeneous in size and shape, are necessary to biofabricate complex tissue and organ models. In addition, when spheroids are produced automatically, there is little chance for microbiological contamination, increasing the reproducibility of the method.
The large-scale production of spheroids is considered the first mandatory step for developing a biofabrication line, which continues in the 3D bioprinting process and finishes in the full maturation of the tissue construct in bioreactors. However, the number of studies that explored the large-scale ASC spheroid production are still scarce, together with the number of studies that used ASC spheroids as building blocks for 3D bioprinting. Therefore, this article aims to show the large-scale production of ASC spheroids using a non-adhesive micromolded hydrogel technique spreading ASC spheroids as building blocks for 3D bioprinting approaches.

Here, we describe the large-scale production of adipose-derived stromal/stem cell (ASC) spheroids using an automated pipetting system to seed the cell suspension, thus ensuring homogeneity of spheroid size and shape. These ASC spheroids can be used as building blocks for 3D bioprinting approaches.

Adipose-derived stromal/stem cells (ASCs) are a subpopulation of cells found in the stromal vascular fraction of human subcutaneous adipose tissue ...

Ensemble Force Spectroscopy by Shear Forces

Single-molecule techniques based on fluorescence and mechanochemical principles provide superior sensitivity in biological sensing. However, due to the lack of high throughput capabilities, the application of these techniques is limited in biophysics. Ensemble force spectroscopy (EFS) has demonstrated high throughput in the investigation of a massive set of molecular structures by converting mechanochemical studies of individual molecules into those of molecular ensembles. In this protocol, the DNA secondary structures (i-motifs) were unfolded in the shear flow between the rotor and stator of a homogenizer tip at shear rates up to 77796/s. The effects of flow rates and molecular sizes on the shear forces experienced by the i-motif were demonstrated. The EFS technique also revealed the binding affinity between DNA i-motifs and ligands. Furthermore, we have demonstrated a click chemistry reaction that can be actuated by shear force (i.e., mechano-click chemistry). These results establish the effectiveness of using shear force to control the conformation of molecular structures.

Ensemble force spectroscopy (EFS) is a robust technique for mechanical unfolding and real-time sensing of an ensemble set of biomolecular structures in biophysical and biosensing fields.

Single-molecule techniques based on fluorescence and mechanochemical principles provide superior sensitivity in biological sensing. However, due to the ...