R.K.B. acknowledges support from the Presidential Postdoctoral Research Fellows Program. This material is also based upon work supported by NSF Graduate Research Fellowship Program Grant DGE-2039656 (to A.M.H.). A.S.D.-M. and H.N.L. acknowledge support from the Lidow Independent Work/Senior Thesis Fund at Princeton University. We also thank the laboratory of Bonnie Bassler for providing strains of V. cholerae. S.S.D. acknowledges support from NSF Grants CBET-1941716, DMR-2011750, and EF-2124863, as well as the Eric and Wendy Schmidt Transformative Technology Fund, the New Jersey Health Foundation, the Pew Biomedical Scholars Program, and the Camille Dreyfus Teacher-Scholar Program.

This approach is to convert a commercial 3D fused deposition modeling printer into a 3D bioprinter using a previously established protocol by Tashman et al.34. In brief, Tashman et al. replaced a commercial extruder head with a custom-made syringe pump extruder. This extruder enables the printing of highly concentrated liquid suspensions of bacterial cells in 3D, with its extruded volume and 3D position controlled by the G-code programming language. The extruded volume is specified in the software by the extruder step (E-step) and is additionally calibrated as described further below. These bacterial suspensions are thereby printed directly into a granular hydrogel matrix, which acts as a 3D support for the cells. Below, the protocol also describes how to prepare matrices with different polymer concentrations, characterize the resulting changes in pore size and rheological properties, and characterize subsequent bacterial motility and growth using direct imaging.
1. Conversion of a commercial 3D printer into a 3D bioprinter
<ol>
	<li>Remove the extruder and the heater from a commercial 3D printer (see Table of Materials).</li>
	<li>Follow previous protocols to fabricate the syringe pump extruder34, with an additional modification to accommodate a disposable Luer lock syringe. Mount the syringe pump extruder onto the printer. 
	NOTE: The CAD files required for modifying the syringe pump extruder for plastic syringes are provided in&#160;Supplementary Files 1-3.</li>
	<li>Install and open the 3D printer software (see Table of Materials) on a computer. Connect the 3D printer to the computer.</li>
	<li>Load a 1 mL disposable syringe with an appropriately sized needle into the 3D-printed clamps by aligning the top and bottom halves of the mechanism (Figure 1). Secure the clamps around the syringe using three M8 socket bolts and thin steel hex nuts (see Table of Materials). The syringe plunger connects with the lead screw in the syringe pump extruder. Manually raise the plunger by rotating the lead screw to create an 0.5 mL of air gap in the syringe.</li>
	<li>If contamination is a concern for the experiment, transport the syringe-clamp complex to a biohood and sterilize with 70% ethanol spray upon entry before completing the following steps.</li>
</ol>
2. Preparation of the bacterial suspension
<ol>
	<li>For V. cholerae and E. coli, grow&#160;overnight on a 2% Lennox LB (Luria Broth, see Table of Materials) agar plate at 37 &#176;C.
	<ol>
		<li>For V. cholerae, inoculate the cells into 3 mL of liquid LB with ten sterile glass beads. Grow the cells in a shaking incubator at 37 &#176;C for 5-6 h until the mid-exponential phase to an optical density (OD) 600 of ~0.9.</li>
		<li>For E. coli, inoculate the cells into 3 mL of liquid LB. Grow the cells in a shaking incubator at 37 &#176;C overnight. Innoculate 200 &#181;L of the overnight culture in fresh LB for 3 h until the OD reaches 0.6.</li>
	</ol>
	</li>
	<li>Transfer the culture into a 10 mL centrifuge tube. Centrifuge the culture for 5 min at 5,000 x g at room temperature to form a pellet. Remove the supernatant. Resuspend with ~10 &#181;L of liquid LB to achieve a cell density of ~9 x 1010 cells per mL.</li>
</ol>
3. Loading the bacterial suspension into the syringe
NOTE: Two methods are provided for loading the bacteria into the syringe (step 3.1 and step 3.2). Step 3.1 works for loading small volumes of bacterial suspensions, &#60;200 &#181;L, and step 3.2 works for loading larger volumes of bacterial suspensions, &#62;200 &#181;L. Step 3.1 was utilized for the representative results shown here.
<ol>
	<li>Load an empty 1 mL plastic Luer lock syringe into the 3D bioprinter. Connect the syringe plunger with the lead screw. Manually retract the syringe to add 0.2 mL of the air gap to provide space for the plunger to move in the syringe as a small volume of cells ~20-50 &#181;L are used for each batch of experiments.
	<ol>
		<li>Attach a blunt needle to the syringe tip with the needle size necessary for the size of print features required. Here, a 2-inch 20 G needle is used.</li>
		<li>Load the bacterial suspension into the syringe by placing a 10 mL centrifuge tube with the bacterial inoculum below the needle. Manually rotate the screw to retract the syringe plunger and load the cells into the syringe. The bacterial cell volumes are so small that often, the cells are only loaded into the needle.</li>
	</ol>
	</li>
	<li>Remove the plunger from the syringe-clamp complex and use another syringe and needle to carefully load the syringe-clamp complex with the desired bacterial suspensions, being careful to avoid trapping air bubbles. The syringe-clamp complex should be filled slightly over the brim with the desired bacterial suspension and then transferred to the bioprinter.
	<ol>
		<li>Carefully insert the syringe-clamp complex without the plunger into the corresponding socket on the main core of the bio-printer extruder.</li>
		<li>Ensure that the printer carriage is approximately halfway up the lead screw, and a collection dish is present under the loaded syringe. Then, carefully insert the plunger through both the carriage and the syringe-clamp complex until it catches on the carriage. Depress the plunger slowly into the bacterial suspension to avoid trapping air bubbles in the syringe.</li>
		<li>Slide the adaptor clamp onto the carriage over the back of the plunger to secure it in place for both extrusion and retraction maneuvers.</li>
	</ol>
	</li>
</ol>
4. Calibration of the extruder step to deposited volume
<ol>
	<li>To calibrate the extruder step (E-step) to the deposited volume, first set up the bio-printer with the exact syringe, syringe needle, and depositing bacterial suspension that will be used in the experiment. Here, a 1 mL Luer lock syringe is used.</li>
	<li>Determine an estimated E-step range to calibrate over by extruding an arbitrary E-step number (~200) and note the plunger volume change with syringe volume markers.</li>
	<li>Use this coarse E-step to volume ratio to determine the E-step settings to perform the calibration sweep over. For example, if an E-step of 200 extrudes approximately 20 &#181;L by visual inspection and one wants to deposit 10-200 &#181;L, test E-steps between 100-2000.</li>
	<li>To perform the linear calibration sweep, first label and measure the dry mass of twenty 1.5 mL sampling tubes on an analytical balance with 0.1 mg sensitivity.</li>
	<li>Extrude bacterial suspension into the pre-measured 1.5 mL tubes. For each E-step, perform at least 2 replicates. Repeat for all E-steps over the linear range, replacing bacterial suspension as necessary. If contamination is a concern for the experiment, wipe the exterior of the syringe needle with a lint-free wipe saturated with 70% ethanol after each sample.</li>
	<li>Measure the mass of all 1.5 mL tubes with the same analytical balance. Subtract the first mass value from the second to obtain a net mass of bacterial suspension deposited.</li>
	<li>Convert the mass of bacterial suspension into a volume with the material density. For many bacterial suspensions composed primarily of water, 1 g/mL is an appropriate density approximation.</li>
	<li>Perform a linear fit between the E-step and the extruded volume to finish the calibration process.</li>
</ol>
5. Preparation of the granular hydrogel matrix
<ol>
	<li>In a biosafety cabinet, add dry granules of cross-linked acrylic acid/alkyl acrylate copolymers (see Table of Materials) to 400 mL of 2% Lennox Luria-Bertani (LB) to keep the matrix sterile; however, other liquid cell culture media can also be used to swell the hydrogel matrix. 
	NOTE: The weight percent of granules added to the LB depends on the pore size that one is aiming for. In the present study, for a 0.9% granular hydrogel matrix, 3.6 g of dry granules are added to the LB and for a 1.2% granular hydrogel matrix, 4.8 g of dry granules are added to the LB. The hydrogel granules are homogenously dispersed by mixing them in a stand mixer for 2 min.</li>
	<li>Once mixed, adjust the pH to 7.4 by adding 20 to 500 &#181;L increments of 10 M sodium hydroxide (NaOH) to ensure cell viability. After each addition of NaOH, measure the pH by dipping a pipette tip into the mixture and then wiping the hydrogel matrix onto a pH test paper. 
	NOTE: As the NaOH is added, the viscosity of the mixture will increase as the hydrogel granules start to swell. The swollen hydrogel granules are ~5 &#181;m to 10 &#181;m in diameter and jammed together in a hydrogel matrix. The internal mesh size of the granules is ~40 nm to 100 nm, as previously established32. The mesh size is large enough for small molecules (e.g., oxygen and nutrients) to freely diffuse but small enough for bacteria to be confined between the interstitial pores.</li>
	<li>Next, transfer the granular hydrogel matrix to a 50 mL centrifuge tube using a 50 mL sterile plastic syringe. Centrifuge the hydrogel matrix at 161 x g for 1 min at room temperature to remove the bubbles formed during the mixing process.</li>
	<li>Allow the hydrogel matrix to sit for at least 2 days at room temperature to ensure that no contamination has occurred. The contamination appears as microcolonies suspended in the hydrogel matrix. After two days, centrifuge the hydrogel matrix at 161 x g for 1 min to remove any additional bubbles that formed. 
	NOTE: The protocol can be paused here by storing the hydrogel matrix at room temperature for up to a week.</li>
	<li>In the biosafety cabinet, using a 30 mL sterile plastic syringe, transfer the desired amount of hydrogel matrix to the container where printing will occur (here ~20 mL for a 20 mL tissue culture flask or 1 mL for 1 mL plastic micro cuvettes were used).</li>
</ol>
6. Characterization of the granular hydrogel matrix rheological properties
<ol>
	<li>Load ~3 mL of the hydrogel matrix into a shear rheometer (see Table of Materials) with a 1 mm gap between roughened 50 mm diameter parallel plates to measure the rheological properties.</li>
	<li>Quantify the yield behavior using unidirectional shear measurements on the shear rheometer by measuring the shear stress as a function of a logarithmic sweep of shear rate from 10-4 s-1 to 102 s-1 (e.g., Figure 2A). 
	NOTE: At low shear rates, the hydrogel matrix will have a constant shear stress (the yield stress) that is independent of the shear rate. At high shear rates, the shear stress will increase with a power-law dependence on the shear rate, indicating the fluidizing of the hydrogel matrix. This yield-stress behavior allows bacteria to be 3D printed within the granular hydrogel matrix29.</li>
	<li>Measure the storage and loss moduli, G&#39; and G&#39;&#39; respectively, as a function of frequency using small amplitude oscillatory rheology with a strain amplitude of 1% and frequencies between 0.1 to 1 Hz (e.g., Figure 2B). 
	NOTE: The ideal granular hydrogel matrix for 3D printing should have a storage modulus larger than the loss modulus, which indicates the medium acts as a jammed elastic solid29.</li>
</ol>
7. Characterization of the granular hydrogel matrix interstitial pore size
<ol>
	<li>Sonicate 100 nm carboxylated fluorescent polystyrene nanoparticles (~3.6 x 1013 particles/mL, see Table of Materials) in their packaging for 15 min to resuspend to break up any aggregations/clusters of particles. Transfer 50 &#181;L of nanoparticles to a 1.5 mL microcentrifuge tube.
	<ol>
		<li>Centrifuge for 10 min at 9,500 x g at room temperature until the pellet forms and the supernatant is clear. Remove the supernatant and resuspend the pellet in 1 mL of the liquid growth media (here LB) used to prepare the granular hydrogel matrix. 
		NOTE: The protocol can be paused here by storing the resuspended nanoparticles at 4 &#176;C for up to 3 months.</li>
	</ol>
	</li>
	<li>Sonicate the resuspended nanoparticles for 30 min. Transfer 1 mL of granular hydrogel matrix to a 1.5 mL microcentrifuge tube. Add 1 &#181;L of the resuspended nanoparticles to the granular hydrogel matrix and mix them with a pipette tip. After mixing, centrifuge for 30 s at 161 x g at room temperature.</li>
	<li>Transfer the hydrogel matrix and the nanoparticle mixture to a 35 mm diameter Petri dish with a 0.1 mm thick glass bottom well. The well is 20 mm in diameter and 1 mm in depth. Place a glass coverslip on top and press down to disable flow and evaporation during imaging. An alternative to the glass coverslip is to add 1 mL of paraffin oil on the top.</li>
	<li>Image the nanoparticles using a confocal microscope with a 40x oil objective with 8x additional zoom in the imaging software (see Table of Materials). 
	NOTE: A higher magnification objective could be used instead of using additional digital zoom in the software.
	<ol>
		<li>Image a time loop with no delay (ideally ~19 frames/s) in a single z-plane for 2 min with at least four nanoparticles within the field of view. Repeat 15 to 20 times in different locations to collect enough data for statistics (100 to 200 nanoparticles).</li>
	</ol>
	</li>
	<li>Use a particle tracking software to analyze the displacement of the particles. Here, a custom-written script is used based on the classic Crocker-Grier algorithm to track the center of mass of the nanoparticle35 (see Supplementary File 7).
	<ol>
		<li>From the particle tracking, calculate the mean square displacement (MSD). The MSD will exhibit free diffusion in the pore space at short lengths and time scales and transition to sub-diffusive scaling at large lengths and time scales due to confinement35.</li>
	</ol>
	</li>
	<li>Identify the length scale where the transition to sub-diffusive scaling occurs to estimate the local pore size. Calculate pore size by adding this length scale to the nanoparticle diameter. Repeat the pore size analysis for every nanoparticle measured. This will yield a pore size distribution from which a mean pore size can be calculated (e.g., Figure 3).</li>
</ol>
8. 3D printing process
<ol>
	<li>3D-print custom-made holders for the sample containers (see Supplementary Files 4-6 for the CAD files). Here, holders for the tissue culture flasks and microcuvettes are used. The holders allow for programming the printer to print multiple samples in a single print session. Place the sample containers with hydrogel media in the holders on the build platform.</li>
	<li>Open the 3D printing software. Load a pre-programmed g-code into the software. For the representative results, step 3.1 is used to load the bacterial suspension into the 3D printer. 
	NOTE: An example of a g-code for printing linear vertical geometries is given in Table 1.</li>
	<li>Through the 3D printing software, move the x-y-z planes to center the print head on the x-y plane to be the first container and then home the z-axis. The homing z-axis will raise the print head. Manually rotate the screw slowly to depress the syringe plunger until a small amount of the bacterial suspension can be seen at the tip of the needle.
	<ol>
		<li>Lightly wipe the excess bacterial suspension off with a sterile disposable wipe. Based on the height of the sample holder, needle, and syringe, using the 3D printing software, lower the print head a fixed distance into the hydrogel media in the sample container of choice. Start the printing process by clicking on Print.</li>
	</ol>
	</li>
	<li>Once the printing is complete, close the sample containers. Wipe down the printer with 70% ethanol. Properly dispose of the syringe and needle.</li>
</ol>
9. Growing and imaging the V. cholerae
<ol>
	<li>For large field-of-view imaging, use a camera with a zoom lens attachment to image cell growth with a lightbox. Image the samples right after printing at room temperature and then transfer them to an incubator. Maintain samples at 37 &#176;C in a stationary incubator between imaging sessions during the experiment.</li>
	<li>Capture images over a desired amount of time to observe the growth behavior at long periods in the granular hydrogel matrix.</li>
</ol>

Bioprinting Bacteria in Hydrogel Matrices for the Study of Motility and Growth

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+semi-solid+agar+for+the+detection+of+bacterial+motility.">The use of semi-solid agar for the detection of bacterial motility.</a> Journal of Bacteriology. 31 (6), 575-580 (1936).</li><li>Bhattacharjee, T., 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=Polyelectrolyte+scaling+laws+for+microgel+yielding+near+jamming.">Polyelectrolyte scaling laws for microgel yielding near jamming.</a> Soft Matter. 14 (9), 1559-1570 (2018).</li><li>Bhattacharjee, T., Datta, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Confinement+and+activity+regulate+bacterial+motion+in+porous+media.">Confinement and activity regulate bacterial motion in porous media.</a> Soft Matter. 15 (48), 9920-9930 (2019).</li><li>Bhattacharjee, T., Datta, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+hopping+and+trapping+in+porous+media.">Bacterial hopping and trapping in porous media.</a> Nature Communications. 10 (1), 2075 (2019).</li><li>Bhattacharjee, T., Amchin, D. B., Ott, J. A., Kratz, F., Datta, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotactic+migration+of+bacteria+in+porous+media.">Chemotactic migration of bacteria in porous media.</a> Biophysical Journal. 120 (16), 3483-3497 (2021).</li><li>Bhattacharjee, T., Amchin, D. B., Alert, R., Ott, J. A., Datta, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotactic+smoothing+of+collective+migration.">Chemotactic smoothing of collective migration.</a> eLife. 11, e71226 (2022).</li><li>Tashman, J. W., Shiwarski, D. J., 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=A+high+performance+open-source+syringe+extruder+optimized+for+extrusion+and+retraction+during+FRESH+3D+bioprinting.">A high performance open-source syringe extruder optimized for extrusion and retraction during FRESH 3D bioprinting.</a> HardwareX. 9, e00170 (2021).</li><li>Crocker, J. C., Grier, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+digital+video+microscopy+for+colloidal+studies.">Methods of digital video microscopy for colloidal studies.</a> Journal of Colloid and Interface Science. 179 (1), 298-310 (1996).</li><li>Chatterjee, T., Chatterjee, B. K., Chakrabarti, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modelling+of+growth+kinetics+of+Vibrio+cholerae+in+presence+of+gold+nanoparticles:+Effect+of+size+and+morphology.">Modelling of growth kinetics of Vibrio cholerae in presence of gold nanoparticles: Effect of size and morphology.</a> Scientific Reports. 7 (1), 9671 (2017).</li><li>Mart&#237;nez-Calvo, 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=Morphological+instability+and+roughening+of+growing+3D+bacterial+colonies.">Morphological instability and roughening of growing 3D bacterial colonies.</a> Proceedings of the National Academy of Sciences. 119 (43), e2208019119 (2022).</li><li>Lehner, B. A. E., Schmieden, D. T., Meyer, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Straightforward+approach+for+3D+bacterial+printing.">A Straightforward approach for 3D bacterial printing.</a> ACS Synthetic Biology. 6 (7), 1124-1130 (2017).</li><li>Zhang, Q., 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=Morphogenesis+and+cell+ordering+in+confined+bacterial+biofilms.">Morphogenesis and cell ordering in confined bacterial biofilms.</a> Proceedings of the National Academy of Sciences. 118 (31), e2107107118 (2021).</li></ol>

The experimental platform used to 3D print and image bacterial communities in this publication is the subject of a patent application filed by Princeton University on behalf of Tapomoy Bhattacharjee and S.S.D. (PCT Application number PCT/US/2020/030213).

Critical steps in the protocol 
It is important to ensure that when preparing each hydrogel matrix, the matrix is made in a sterile environment. If not, contamination can occur, which manifests as, e.g., microcolonies (small spheroids) in the matrix after several days. During the mixing process, it is important that all the dry granular hydrogel particles are dissolved. Additionally, when adjusting the pH of each hydrogel matrix with the NaOH, the granules will start to swell, which increases the viscosity of the hydrogel matrix, leading to mixing being more difficult. Using the stand mixer will help ensure that the NaOH is well mixed into the hydrogel matrix. During the loading of each bacterial suspension, air pockets can form in the needle. To avoid this issue, ensure that the needle tip is always sitting in the bacterial suspension in the centrifuge tube and not at the bottom of the tube or near the top surface. Another way to overcome this issue is to grow large volumes of cells and thus have larger volumes of the bacterial suspension for printing.
Limitations 
Currently, during printing, the low viscosity of the bacterial suspension limits the geometries that can be printed and often leads to a biofilm-forming and growing on the top of the hydrogel matrix surface due to trace cells. There are a few potential methods for overcoming this limitation, including increasing the viscosity of the bacterial suspension or further optimizing the 3D printer settings. To increase the viscosity of the bacterial suspension, one could mix the bacterial suspension with another polymer - for example, alginate, which has been used prior for the 3D printing of bacteria onto flat surfaces38. The printer settings can be further optimized to enable retraction of the syringe plunger during the withdrawal of the needle from the granular hydrogel matrix, which would have the potential to stop cells from being deposited during the removal of the needle from the hydrogel matrix.
The significance of the method with respect to existing/alternative methods 
The method described here allows for the printing of bacterial colonies into granular hydrogel matrices. The granular hydrogel matrices allow the study of the impact of external environmental factors (e.g., pore size, matrix deformability) on the motility and growth of bacteria. Additionally, while in this work, LB is used as the liquid growth medium to swell the hydrogel matrix, the hydrogel matrix can be swollen with other liquid growth media, including media with antibiotics. Previous methods for studying bacteria in confined environments were limited by the length of experimental time, the polymer mesh size, and surrounding hydrogel matrix stiffness37,38. Protocols already exist for making granular hydrogel matrices out of different polymers, so the potential for studying the impacts of different environmental conditions on the motility and growth of bacteria is vast. This method allows for the study of bacteria in control environments that more readily recapitulate the environments that bacteria inhabit in the real world, such as host mucus or soil. Another limitation of many other methods is the opacity of the surrounding matrix; however, this approach using optically transparent materials provides the ability to explore, e.g., optogenetic control and patterning of bacteria in 3D.
Beyond studying motility and growth, the 3D printing method described here overcomes the limitation of many other bioprinting methods that require the deposition of a bioink on a substrate and are, therefore, limited in the height of the engineered living material they can produce. In the future, this bioprinting protocol can be further expanded to fabricate biohybrid materials by mixing polymers with biofilm-forming cells. The granular hydrogel matrices provide support for 3D printing thicker, larger-scale engineered living materials and more complex geometries than many other current bacteria bioprinting methods. While this work only used V. cholerae and E. coli, other species, such as Pseudomonas aeruginosa, have also successfully been 3D printed37. Beyond printing, the printer can be adapted to do a controlled sampling of bacteria after growth to see if there have been any genetic changes, for example.

Bacteria often inhabit diverse, complex 3D porous environments ranging from mucosal gels in the gut and lungs to soil in the ground1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18,19,20,21, 
22,23,24,25. In these settings, bacterial movement through motility or growth can be impeded by surrounding obstacles, such as polymer networks or packings of solid mineral grains-influencing the ability of the cells to spread through their environments26, access nutrient sources, colonize new terrain, and form protective biofilm communities27. However, traditional lab studies typically employ highly simplified geometries, focusing on cells in liquid cultures or on flat surfaces. While these approaches yield key insights into microbiology, they do not fully recapitulate the complexity of natural habitats, leading to dramatic differences in growth rates and motility behavior compared to measurements performed in real-world settings. Therefore, a method to define bacterial colonies and study their motility and growth in 3D porous environments more akin to many of their natural habitats is critically needed.
Inoculating cells into an agar gel and then visualizing their macroscopic spreading by eye or using a camera provides one straightforward way to accomplish this, as first proposed by Tittsler and Sandholzer in 193628. However, this approach suffers from a number of key technical challenges: (1) While the pore sizes can, in principle, be varied by varying the agarose concentration, the pore structure of such gels is poorly defined; (2) Light scattering causes these gels to be turbid, making it difficult to visualize cells at the individual scale with high resolution and fidelity, particularly in large samples; (3) When the agar concentration is too large, cell migration is restricted to the top flat surface of the gel; (4) The complex rheology of such gels makes it challenging to introduce inocula with well-defined geometries.
To address these limitations, in previous work, Datta&#39;s lab developed an alternate approach using granular hydrogel matrices - comprised of jammed, biocompatible hydrogel particles swollen in liquid bacterial culture - as &#34;porous Petri dishes&#34; to confine cells in 3D. These matrices are soft, self-healing, yield-stress solids; thus, unlike with cross-linked gels used in other bioprinting processes, an injection micronozzle can move freely inside the matrix along any prescribed 3D path by locally rearranging the hydrogel particles29. These particles then re-densify rapidly and self-heal around injected bacteria, supporting the cells in place without any additional harmful processing. This process is, therefore, a form of 3D printing that enables bacterial cells to be arranged - in a desired 3D structure, with a defined community composition - within a porous matrix having tunable physicochemical properties. Moreover, the hydrogel matrices are completely transparent, enabling the cells to be directly visualized using imaging.
The utility of this approach has been demonstrated previously in two ways. In one set of studies, dilute cells were dispersed throughout the hydrogel matrix, which enabled studies of the motility of individual bacteria30,31. In another set of studies, multicellular communities were 3D-printed in centimeter-scale gels using an injection nozzle mounted on a programmable microscope stage, which enabled studies of the spreading of bacterial collectives through their surroundings32,33. In both cases, these studies revealed previously unknown differences in the spreading characteristics of bacteria inhabiting porous environments compared to those in liquid culture/on flat surfaces. However, given that they were mounted on a microscope stage, these previous studies were limited to small sample volumes (~1 mL) and, therefore, short experimental time scales. They were also limited in their ability to define inocula geometries with high spatial resolution.
Here, the next generation of this experimental platform that addresses both limitations is described. Specifically, protocols are provided by which one can use a modified 3D printer with an attached syringe extruder to 3D print and image bacterial colonies at large scales. Moreover, representative data indicates how this approach can be useful for studying the motility and growth of bacteria, using the biofilm-former Vibrio cholerae and planktonic Escherichia coli as examples. This approach enables bacterial colonies to be sustained over long times and visualized using various imaging techniques. Hence, the ability of this approach to study bacterial communities in 3D porous habitats has tremendous research and applied potential, impacting the treatment and study of microbes in the gut, the skin, the lung, and the soil. Moreover, this approach could be used in the future for 3D printing bacteria-based engineered living materials into more complex freestanding shapes.

<table><tbody><tr><td>1 mL cuvettes</td><td>VWR</td><td>97000-586</td><td></td></tr><tr><td>1 mL Luer lock syringe</td><td>BH Supplies</td><td>BH1LL</td><td></td></tr><tr><td>10 M NaOH</td><td>Sigma-Aldrich</td><td>72068</td><td></td></tr><tr><td>100 nm carboxylated fluorescent polystyrene nanoparticles (FluoSpheres)&amp;nbsp;</td><td>&amp;nbsp;Invitrogen, (ThermoFischer Scientific)</td><td>&lt;br/&gt; F8803</td><td></td></tr><tr><td>15 mL centrifuge tubes</td><td>ThermoFischer Scientific</td><td>14-955-237</td><td></td></tr><tr><td>20 G blunt needle</td><td>McMaster Carr</td><td>75165A252</td><td></td></tr><tr><td>25 mL tissue culture flasks</td><td>VWR</td><td>10861-566</td><td></td></tr><tr><td>3D printer</td><td>Lulzbot</td><td>LulzBot Mini 2</td><td></td></tr><tr><td>3D printing software</td><td>Cura</td><td>Cura-Lulzbot</td><td></td></tr><tr><td>50 mL centrifuge tubes</td><td>ThermoFischer Scientific</td><td>14-955-239</td><td></td></tr><tr><td>Agar</td><td>Sigma-Aldrich</td><td>A1296</td><td></td></tr><tr><td>Carbomer Granular Hydrogel Particles</td><td>Lubrizol&amp;nbsp;</td><td>Carbopol 980NF</td><td>dry granules of crosslinked acrylic acid/alkyl acrylate copolymers</td></tr><tr><td>Centrifuge (2 mL tube capacity)</td><td>VWR</td><td>2405-37</td><td></td></tr><tr><td>Centrifuge (50 mL tube capacity)</td><td>ThermoFischer Scientific</td><td>75007200</td><td>Sorvall (brand) ST 8 (model)</td></tr><tr><td>Confocal Microscope</td><td>Nikon</td><td>A1R+ inverted laserscanning&lt;br/&gt; confocal microscope</td><td></td></tr><tr><td>Glass bottom petri dish</td><td>Cellvis</td><td>D35-10-1-N</td><td></td></tr><tr><td>Lennox LB (Lubria Broth)</td><td>Sigma-Aldrich</td><td>L3022</td><td></td></tr><tr><td>M8 &amp;times; 1.25 mm, 150 mm long, Fully Threaded Socket Cap</td><td>McMaster Carr</td><td>91290A478</td><td></td></tr><tr><td>M8 &amp;times; 1.25 mm, Brass Thin Hex Nut</td><td>McMaster Carr</td><td>93187A300</td><td></td></tr><tr><td>Open-source syringe pump</td><td>Custom-made</td><td>Replistruder 4</td><td>https://www.sciencedirect.com/science/article/pii/S2468067220300791</td></tr><tr><td>Petri dish (60 mm round)</td><td>ThermoFischer Scientific</td><td>FB0875713A</td><td></td></tr><tr><td>Shear Rheometer</td><td>Anton Paar</td><td>MCR 501</td><td></td></tr><tr><td>Ultrasonic cleaner</td><td>VWR</td><td>97043-992</td><td></td></tr></tbody></table>

3d printing bacteria to study motility and growth in complex 3d porous media

Bacteria are ubiquitous in complex three-dimensional (3D) porous environments, such as biological tissues and gels, and subsurface soils and sediments. However, the majority of previous work has focused on studies of cells in bulk liquids or at flat surfaces, which do not fully recapitulate the complexity of many natural bacterial habitats. Here, this gap in knowledge is addressed by describing the development of a method to 3D-print dense colonies of bacteria into jammed granular hydrogel matrices. These matrices have tunable pore sizes and mechanical properties; they physically confine the cells, thus supporting them in 3D. They are optically transparent, allowing for direct visualization of bacterial spreading through their surroundings using imaging. As a proof of this principle, here, the capability of this protocol is demonstrated by 3D printing and imaging non-motile and motile Vibro cholerae, as well as non-motile Escherichia coli, in jammed granular hydrogel matrices with varying interstitial pore sizes.

Watch this Scientific Journal Video about Author Spotlight: Studying Bacterial Growth in 3D Hydrogel Matrices at JoVE.com

Author Spotlight: Studying Bacterial Growth in 3D Hydrogel Matrices

This protocol describes a procedure for three-dimensional (3D) printing of bacterial colonies to study their motility and growth in complex 3D porous hydrogel matrices that are more akin to their natural habitats than conventional liquid cultures or Petri dishes.

3D Printing Bacteria to Study Motility and Growth in Complex 3D Porous Media

Bacteria are ubiquitous in complex three-dimensional (3D) porous environments, such as biological tissues and gels, and subsurface soils and sediments. ...

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Research

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Bioengineering

2.1K Views.  University of Colorado Boulder. This protocol describes a procedure for three-dimensional (3D) printing of bacterial colonies to study their motility and growth in complex 3D porous hydrogel matrices that are more akin to their natural habitats than conventional liquid cultures or Petri dishes. 

Video: Author Spotlight: Studying Bacterial Growth in 3D Hydrogel Matrices

From Voxels to Knowledge: A Practical Guide to the Segmentation of Complex Electron Microscopy 3D-Data

Modern 3D electron microscopy approaches have recently allowed unprecedented insight into the 3D ultrastructural organization of cells and tissues, enabling the visualization of large macromolecular machines, such as adhesion complexes, as well as higher-order structures, such as the cytoskeleton and cellular organelles in their respective cell and tissue context. Given the inherent complexity of cellular volumes, it is essential to first extract the features of interest in order to allow visualization, quantification, and therefore comprehension of their 3D organization. Each data set is defined by distinct characteristics, e.g., signal-to-noise ratio, crispness (sharpness) of the data, heterogeneity of its features, crowdedness of features, presence or absence of characteristic shapes that allow for easy identification, and the percentage of the entire volume that a specific region of interest occupies. All these characteristics need to be considered when deciding on which approach to take for segmentation.
The six different 3D ultrastructural data sets presented were obtained by three different imaging approaches: resin embedded stained electron tomography, focused ion beam- and serial block face- scanning electron microscopy (FIB-SEM, SBF-SEM) of mildly stained and heavily stained samples, respectively. For these data sets, four different segmentation approaches have been applied: (1) fully manual model building followed solely by visualization of the model, (2) manual tracing segmentation of the data followed by surface rendering, (3) semi-automated approaches followed by surface rendering, or (4) automated custom-designed segmentation algorithms followed by surface rendering and quantitative analysis. Depending on the combination of data set characteristics, it was found that typically one of these four categorical approaches outperforms the others, but depending on the exact sequence of criteria, more than one approach may be successful. Based on these data, we propose a triage scheme that categorizes both objective data set characteristics and subjective personal criteria for the analysis of the different data sets.

The bottleneck for cellular 3D electron microscopy is feature extraction (segmentation) in highly complex 3D density maps. We have developed a set of criteria, which provides guidance regarding which segmentation approach (manual, semi-automated, or automated) is best suited for different data types, thus providing a starting point for effective segmentation.

Modern 3D electron microscopy approaches have recently allowed unprecedented insight into the 3D ultrastructural organization of cells and tissues, ...

Combining Fluidic Devices with Microscopy and Flow Cytometry to Study Microbial Transport in Porous Media Across Spatial Scales

Understanding the transport, dispersion and deposition of microorganisms in porous media is a complex scientific task comprising topics as diverse as hydrodynamics, ecology and environmental engineering. Modeling bacterial transport in porous environments at different spatial scales is critical to better predict the consequences of bacterial transport, yet current models often fail to up-scale from laboratory to field conditions. Here, we introduce experimental tools to study bacterial transport in porous media at two spatial scales. The aim of these tools is to obtain macroscopic observables (such as breakthrough curves or deposition profiles) of bacteria injected into transparent porous matrices. At the small scale (10-1000 &#181;m), microfluidic devices are combined with optical video-microscopy and image processing to obtain breakthrough curves and, at the same time, to track individual bacterial cells at the pore scale. At larger scale, flow cytometry is combined with a self-made robotic dispenser to obtain breakthrough curves. We illustrate the utility of these tools to better understand how bacteria are transported in complex porous media such as the hyporheic zone of streams. As these tools provide simultaneous measurements across scales, they pave the way for mechanism-based models, critically important for upscaling. Application of these tools may not only contribute to the development of novel bioremediation applications but also shed new light on the ecological strategies of microorganisms colonizing porous substrates.

Breakthrough curves (BTCs) are efficient tools to study the transport of bacteria in porous media. Here we introduce tools based on fluidic devices in combination with microscopy and flow cytometric counting to obtain BTCs.

Understanding the transport, dispersion and deposition of microorganisms in porous media is a complex scientific task comprising topics as diverse as ...

Environment

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 ...

2D and 3D Matrices to Study Linear Invadosome Formation and Activity

Cell adhesion, migration, and invasion are involved in many physiological and pathological processes. For example, during metastasis formation, tumor cells have to cross anatomical barriers to invade and migrate through the surrounding tissue in order to reach blood or lymphatic vessels. This requires the interaction between cells and the extracellular matrix (ECM). At the cellular level, many cells, including the majority of cancer cells, are able to form invadosomes, which are F-actin-based structures capable of degrading ECM. Invadosomes are protrusive actin structures that recruit and activate matrix metalloproteinases (MMPs). The molecular composition, density, organization, and stiffness of the ECM are crucial in regulating invadosome formation and activation. In vitro, a gelatin assay is the standard assay used to observe and quantify invadosome degradation activity. However, gelatin, which is denatured collagen I, is not a physiological matrix element. A novel assay using type I collagen fibrils was developed and used to demonstrate that this physiological matrix is a potent inducer of invadosomes. Invadosomes that form along the collagen fibrils are known as linear invadosomes due to their linear organization on the fibers. Moreover, molecular analysis of linear invadosomes showed that the discoidin domain receptor 1 (DDR1) is the receptor involved in their formation. These data clearly demonstrate the importance of using a physiologically relevant matrix in order to understand the complex interactions between cells and the ECM.

This protocol describes how to prepare a 2D mixed matrix, consisting of gelatin and collagen I, and a 3D collagen I plug to study linear invadosomes. These protocols allow for the study of linear invadosome formation, matrix degradation activity, and the invasion capabilities of primary cells and cancer cell lines.

Cell adhesion, migration, and invasion are involved in many physiological and pathological processes. For example, during metastasis formation, tumor ...

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 ...

Novel Process for 3D Printing Decellularized Matrices

3D bioprinting aims to create custom scaffolds that are biologically active and accommodate the desired size and geometry. A thermoplastic backbone can provide mechanical stability similar to native tissue while biologic agents offer compositional cues to progenitor cells, leading to their migration, proliferation, and differentiation to reconstitute the original tissues/organs1,2. Unfortunately, many 3D printing compatible, bioresorbable polymers (such as polylactic acid, PLA) are printed at temperatures of 210 &#176;C or higher - temperatures that are detrimental to biologics. On the other hand, polycaprolactone (PCL), a different type of polyester, is a bioresorbable, 3D printable material that has a gentler printing temperature of 65 &#176;C. Therefore, it was hypothesized that decellularized extracellular matrix (DM) contained within a thermally protective PLA barrier could be printed within PCL filament and remain in its functional conformation. In this work, osteochondral repair was the application for which the hypothesis was tested. As such, porcine cartilage was decellularized and encapsulated in polylactic acid (PLA) microspheres which were then extruded with polycaprolactone (PCL) into filament to produce 3D constructs via fused deposition modeling. The constructs with or without the microspheres (PLA-DM/PCL and PCL(-), respectively) were evaluated for differences in surface features.

This protocol describes the production of polycaprolactone (PCL) filament with embedded polylactic acid (PLA) microspheres which contain decellularized matrices (DM) for 3D printing of structural tissue engineering constructs.

3D bioprinting aims to create custom scaffolds that are biologically active and accommodate the desired size and geometry. A thermoplastic backbone can ...

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and chemical complexity of living tissues and mimic in vivo tissue functions. Recent advances in microfabrication technologies have facilitated the development of 3D in vitro environments in which cells can be integrated into a well-defined extracellular matrix (ECM) and a defined set of soluble or matrix associated biomolecules. However, technological barriers have limited their widespread use in research laboratories. Here, we describe a method to construct simple devices for 3D culture and experimentation with cells and multicellular organoids in 3D microenvironments with a defined chemoattractant gradient. We illustrate the use of this platform for analysis of the response of epithelial cells and organoids to gradients of growth factors, such as epidermal growth factor (EGF). EGF gradients were stable in the devices for several days leading to directed branch formation in breast organoids. This analysis allowed us to conclude that collective gradient sensing by groups of cells is more sensitive vs. single cells. We also describe the fabrication method, which does not require photolithography facilities nor advanced soft lithography techniques. This method will be helpful to study 3D cellular behaviors in the context of the analysis of development and pathological states, including cancer.

We describe a method to construct devices for 3D culture and experimentation with cells and multicellular organoids. This device allows analysis of cellular responses to soluble signals in 3D microenvironments with defined chemoattractant gradients. Organoids are better than single cells at detection of weak noisy inputs.

Various limitations of 2D cell culture systems have sparked interest in 3D cell culture and analysis platforms, which would better mimic the spatial and ...

Three-dimensional Patterning of Engineered Biofilms with a Do-it-yourself Bioprinter

Biofilms are aggregates of bacteria embedded in a self-produced spatially-patterned extracellular matrix. Bacteria within a biofilm develop enhanced antibiotic resistance, which poses potential health dangers, but can also be beneficial for environmental applications such as purification of drinking water. The further development of anti-bacterial therapeutics and biofilm-inspired applications will require the development of reproducible, engineerable methods for biofilm creation. Recently, a novel method of biofilm preparation using a modified three-dimensional (3D) printer with a bacterial ink has been developed. This article describes the steps necessary to build this efficient, low-cost 3D bioprinter that offers multiple applications in bacterially-induced materials processing. The protocol begins with an adapted commercial 3D printer in which the extruder has been replaced with a bio-ink dispenser connected to a syringe pump system enabling a controllable, continuous flow of bio-ink. To develop a bio-ink suitable for biofilm printing, engineered Escherichia coli bacteria were suspended in a solution of alginate, so that they solidify in contact with a surface containing calcium. The inclusion of an inducer chemical within the printing substrate drives expression of biofilm proteins within the printed bio-ink. This method enables 3D printing of various spatial patterns composed of discrete layers of printed biofilms. Such spatially-controlled biofilms can serve as model systems and can find applications in multiple fields that have a wide-ranging impact on society, including antibiotic resistance prevention or drinking water purification, among others.

This article describes a method of transforming a low-cost commercial 3D printer into a bacterial 3D printer that can facilitate printing of patterned biofilms. All necessary aspects of preparing the bioprinter and bio-ink are described, as well as verification methods to assess the formation of biofilms.

Biofilms are aggregates of bacteria embedded in a self-produced spatially-patterned extracellular matrix. Bacteria within a biofilm develop enhanced ...

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the microscale&#8213;to create micro-environments for microbial chemotaxis experiments. To create chemical pulses, we developed a system to introduce amino acid sources near-instantaneously by photolysis of caged amino acids within a polydimethylsiloxane (PDMS) microfluidic chamber containing a bacterial suspension. We applied this method to the chemotactic bacterium, Vibrio ordalii, which can actively climb these dynamic chemical gradients while being tracked by video microscopy. Amino acids, rendered biologically inert (&#39;caged&#39;) by chemical modification with a photoremovable protecting group, are uniformly present in the suspension but not available for consumption until their sudden release, which occurs at user-defined points in time and space by means of a near-UV-A focused LED beam. The number of molecules released in the pulse can be determined by a calibration relationship between exposure time and uncaging fraction, where the absorption spectrum after photolysis is characterized by using UV-Vis spectroscopy. A nanoporous polycarbonate (PCTE) membrane can be integrated into the microfluidic device to allow the continuous removal by flow of the uncaged compounds and the spent media. A strong, irreversible bond between the PCTE membrane and the PDMS microfluidic structure is achieved by coating the membrane with a solution of 3-aminopropyltriethoxysilane (APTES) followed by plasma activation of the surfaces to be bonded. A computer-controlled system can generate user-defined sequences of pulses at different locations and with different intensities, so as to create resource landscapes with prescribed spatial and temporal variability. In each chemical landscape, the dynamics of bacterial movement at the individual scale and their accumulation at the population level can be obtained, thereby allowing the quantification of chemotactic performance and its effects on bacterial aggregations in ecologically relevant environments.

A protocol for the generation of dynamic chemical landscapes by photolysis within microfluidic and millifluidic setups is presented. This methodology is suitable to study diverse biological processes, including the motile behavior, nutrient uptake, or adaptation to chemicals of microorganisms, both at the single cell and population level.

We demonstrate a method for the generation of controlled, dynamic chemical pulses&#8213;where localized chemoattractant becomes suddenly available at the ...

Measuring the Migration and Biofilm Formation of Various Bacteria

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their nature would be beneficial in facilitating their characterization. Currently, most studies focus only on one specific characterization method; however, the isolation and identification of microorganisms from the host is a continuous process and usually requires several combinatorial characterization methods. Herein, we describe methods of identifying the microbial biofilm-forming ability, the microbial respiration state, and their chemotaxis behavior. The methods are used to identify five microbes, three of which were isolated from the bone tissue of Sprague-Dawley (SD) rats (Corynebacterium stationis, Staphylococcus cohnii subsp. urealyticus, and Enterococcus faecalis) and two from the American Type Culture Collection (ATCC)-Staphylococcus aureus ATCC 25923&#160;and Enterococcus faecalis V583. The microbes isolated from the SD rat bone tissue include the gram-positive microbes. These microbes have adapted to thrive under stressful and nutrient-limiting environments within the bone matrix. This article aims to provide the readers with the specific know-how of determining the nature and behavior of the isolated microbes within a laboratory setting.

Here, we present a practical method for the isolation and identification of microorganisms within the host. In this way, the physicochemical properties of microorganisms and possible ways of living in the host are clearly described.

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their ...

3D Printing Bacteria to Study Motility and Growth in Complex 3D Porous Media

Summary

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Chapters in this video

From Voxels to Knowledge: A Practical Guide to the Segmentation of Complex Electron Microscopy 3D-Data

Combining Fluidic Devices with Microscopy and Flow Cytometry to Study Microbial Transport in Porous Media Across Spatial Scales

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

2D and 3D Matrices to Study Linear Invadosome Formation and Activity

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

Novel Process for 3D Printing Decellularized Matrices

3D Analysis of Multi-cellular Responses to Chemoattractant Gradients

Three-dimensional Patterning of Engineered Biofilms with a Do-it-yourself Bioprinter

Generating Controlled, Dynamic Chemical Landscapes to Study Microbial Behavior

Measuring the Migration and Biofilm Formation of Various Bacteria