Preparation and use of cellular reagents: a low-resource molecular biology reagent platform

Abstract

Protein reagents are indispensable for most molecular and synthetic biology procedures. Most conventional protocols rely on highly purified protein reagents that require considerable expertise, time, and infrastructure to produce. In consequence, most proteins are acquired from commercial sources, reagent expense is often high, and accessibility might be hampered by shipping delays, customs barriers, geopolitical constraints, and the need for a constant cold chain. Such limitations to the widespread availability of protein reagents, in turn, limit the expansion and adoption of molecular biology methods in research, education, and technology development and application. Here, we describe protocols for producing a low-resource and locally sustainable reagent delivery system, termed ‘cellular reagents’, in which bacteria engineered to overexpress proteins of interest are dried and can then be used in numerous molecular biology reactions directly as reagent packets, without the need for protein purification or a constant cold chain. As an example of their application, we describe the execution of polymerase chain reaction (PCR) and loop-mediated isothermal amplification (LAMP) using cellular reagents, detailing how to replace pure protein reagents with optimal amounts of rehydrated cellular reagents. We additionally describe a DIY fluorescence visualization device for using these cellular reagents in common molecular biology applications. The methods presented in this article can be used for low cost, on-site production of commonly used molecular biology reagents (including DNA and RNA polymerases, reverse transcriptases, and ligases) with minimal instrumentation and expertise, and without requiring protein purification. Consequently, these methods should generally make molecular biology reagents more affordable and accessible.

Basic Protocol 1: Preparation of Cellular Reagents

Alternate Protocol 1: Preparation of lyophilized cellular reagents

Alternate Protocol 2: Evaluation of bacterial culture growth via comparison to McFarland turbidity standards

Support Protocol 1: SDS-PAGE for protein expression analysis of cellular reagents

Basic Protocol 2: Using Taq DNA polymerase cellular reagents for PCR

Basic Protocol 3: Using Br512 DNA polymerase cellular reagents for LAMP

Support Protocol 2: Building a fluorescence visualization device

INTRODUCTION:

Molecular biology procedures such as PCR, reverse transcription, isothermal amplification, and DNA cleavage and assembly are routinely employed both in academic research and in applied settings, including molecular diagnostics. Proteins such as DNA and RNA polymerases, restriction enzymes, and ligases are essential for executing many of these procedures. Most conventional protocols for performing these assays rely on purified reagents and traditional protein purification processes and, therefore, require substantial investments of time, expertise, equipment, and infrastructure (Wingfield, 2015), with a concomitant requirement that most users obtain reagents from commercial sources. Commercial consumable reagent costs are often high and might be encumbered by a need for a constant cold chain (often at below freezing temperatures), long distance shipping delays, and customs and geopolitical barriers. As a result, both affordability and accessibility of protein reagents can be significantly limited, especially in resource-limited or remote field settings (Abou Tayoun, Burchard, Malik, Scherer, & Tsongalis, 2014; Adebamowo et al., 2018). Ultimately, these barriers hamper dissemination and adoption of molecular and synthetic biology techniques for research and education, and for technology development and deployment.

Simplification of production processes, and the elimination of a cold chain during transportation and storage of protein reagents could potentially reduce the cost, time, expertise, and infrastructure needed for reagent manufacture and application and, thereby, increase accessibility and encourage on-site or local production at a less-than-industrial scale. To facilitate such global affordability and unfettered access to molecular biology reagents, we have developed ‘cellular reagents,’ a reagent delivery platform comprised of dried bacteria, such as Escherichia coli K-12, engineered to overexpress proteins of interest. These cellular reagents, which can be used directly to carry out numerous molecular biology reactions (see below), do not require protein purification or a cold chain. As such, they are considerably simpler and cheaper to produce compared with purified or commercial enzymes (Bhadra, Nguyen, et al., 2021).

Preparation of cellular reagents requires only a 37 °C bacterial incubator to grow the protein-expressing bacteria, which are subsequently collected using a tabletop microcentrifuge and then dried by simple overnight incubation in the presence of inexpensive chemical desiccants, such as calcium sulfate. We have also developed standardized protocols for using these cellular reagents directly as bacterial “reagent packets” in lieu of purified proteins for routine procedures for nucleic acid synthesis, assembly, and detection (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018). In such protocols, one can simply replace pure protein reagents with an optimal amount of the corresponding rehydrated cellular reagent, without perturbing other components or reaction parameters.

In this article, we describe general methods for the preparation of cellular reagents and their use in some common molecular biology assays. In Basic Protocol 1, we describe the production of cellular reagents by drying protein-expressing bacteria in the presence of chemical desiccants (Basic Protocol 1). In Alternate Protocol 1, we describe the preparation of cellular reagents by freeze-drying protein-expressing bacteria using a lyophilizer. This protocol is useful when live bacteria must be eliminated from cellular reagent preparations of thermolabile proteins, given that heat-killing, which is typically how live bacteria is eliminated from preparations with thermostable proteins, would be unsuitable. The preparation of cellular reagents in Basic Protocol 1 and Alternate Protocol 1 requires monitoring bacterial growth, and this is done with a spectrophotometer. In Alternate Protocol 2, we detail the assessment of bacterial culture density by comparing its turbidity to McFarland turbidity standards, a method that can be used when a spectrophotometer is not available. Further, in Support Protocol 1, we provide a method for validating protein expression in cellular reagents, by lysing the reagents and analyzing the proteins using sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE).

In addition, we provide protocols describing some applications of cellular reagents. Specifically, we describe the use of Taq DNA polymerase cellular reagents in qPCR (Basic Protocol 2) and engineered Br512 DNA polymerase cellular reagents in loop-mediated isothermal amplification (LAMP) reactions (Basic Protocol 3). Finally, in Support Protocol 2, we describe to the steps to build a fluorescence visualization device, which can be used for readout of LAMP assays in the absence of other devices to detect fluorescence, such as fluorimeters or transilluminators. Figure 1 provides an overview of the protocols described here for the production and use of cellular reagents.

Figure 1. Overview of the protocols described in this article for the production and use of cellular reagents.

Basic Protocol steps where Alternate or Support Protocols are applicable are color coded with matching boxes.

CAUTION: Follow all appropriate aseptic bacterial culture techniques and institutional biosafety guidelines and regulations (Burnett, Lunn, & Coico, 2009; Tuttle, Trahan, & Son, 2021) for preparation, handling, and disposal of bacterial cultures or their byproducts.

STRATEGIC PLANNING

Depending on the type of protein expression construct and intended use of the cellular reagent, prepare or purchase competent cells for the appropriate strain of Escherichia coli and clone your protein of interest in your vector of choice. For instance, BL21(DE3)-based E. coli strains may be used for protein expression constructs relying on T7 RNA polymerase mediated-transcription, while proteins being expressed using E. coli RNA polymerases may be produced in the BL21 E. coli strain. For downstream applications, such as DNA assembly, that might be disrupted by nuclease activity or recombination, E. coli strains bearing endA and recA mutations (e.g., DH5α) may be used to prepare cellular reagents. Typical protocols for preparation and transformation of competent E. coli may be found in (Renzette, 2011). Ready-to-use competent cells and associated transformation protocols are also available from commercial sources, such as New England Biolabs (Ipswich, MA, USA), Promega (Madison, WI, USA), and Thermo Fisher Scientific (Waltham, MA, USA).

The protocol requires users to utilize optimal conditions for bacterial growth and recombinant protein induction. Transform the appropriate bacterial strain with your protein expression construct and test protein induction. If using a previously reported plasmid, utilize the published optimal bacterial growth and protein induction conditions. If working with a new construct, you will need to determine optimal conditions empirically before embarking on the protocol. This can be done by systematically varying parameters such as bacterial density at the time of induction (induce cultures at A₆₀₀ values ranging from 0.5 to 1.0), amount of inducer added to the culture, and temperature and duration of induction (3–4 h at 37 °C to overnight at 18 °C to 30 °C). In each case, verify expression of your protein in induced bacterial cultures by comparing with proteins expressed in duplicate cultures of untransformed bacteria and uninduced transformants, using techniques such as SDS-PAGE or others. In depth guidance on optimizing heterologous protein expression in E. coli may be found in (Francis & Page, 2010).

BASIC PROTOCOL 1: PREPARATION OF CELLULAR REAGENTS

Similar to the preparation of purified recombinant proteins from E. coli (Wingfield, 2014), the preparation of cellular reagents also begins with overexpression of the protein of interest from a desired plasmid in a suitable E. coli strain (LaVallie, 2001; Rosano & Ceccarelli, 2014). Reagents and methods used in this step, such as recombinant protein expression plasmids, bacterial strains and their growth media, transformant selection and plasmid maintenance, and transcription induction procedures, are usually the same as those employed to overexpress recombinant proteins for purification. In a typical preparation, competent bacteria are transformed and plated on selective agar plates for overnight growth. The next day, individual transformants are cultured in liquid bacterial growth medium containing antibiotics, and the following day, a sub-culture is prepared. This is then followed by protein induction under previously optimized conditions (see Strategic Planning). Upon completion of protein production, bacteria are collected by centrifugation, washed, and then dried in aliquots by overnight incubation at 37 °C in the presence of the chemical desiccant calcium sulfate or silica gel. After this incubation, the dry cellular reagents are removed from 37 °C and are ready for use and/or storage. Some of the most commonly used molecular biology enzymes, including DNA and RNA polymerases, reverse transcriptases, and ligases, can be prepared as cellular reagents using this protocol, and should be stable at ambient temperatures for at least several months.

Materials:

• Protein expression plasmid (built de novo or obtained from repositories such as addgene.org or stanford.freegenes.org, or collaboration networks such as reclone.org), 10 to 100 ng/μL

• Competent E. coli strain (BL21 (DE3), New England Biolabs, cat. No. C2527, or equivalent competent cells for protein expression, see Strategic Planning)

• SOC medium (see Reagents and Solutions)

• Superior Broth^™ (Athena Enzyme Systems, cat. No. 0105; or equivalent rich medium (Elbing & Brent, 2019) – see example recipe in Reagents and Solutions)

• Lysogeny Broth (LB) Agar (or equivalent growth medium with agar – see Reagents and Solutions)

• Antibiotics (or equivalent reagents for plasmid maintenance)

• Chemical inducer for protein expression

  • Isopropyl β-d-1-thiogalactopyranoside (IPTG, Sigma-Aldrich, cat. No. I6758, or equivalent reagents for inducing protein expression)
  • Anhydrotetracycline (Caymen Chemicals, cat. No. 10009542, or equivalent reagents for inducing protein expression; see recipes for preparing IPTG and anhydrotetracycline solutions in Reagents and Solutions)

• 1X Phosphate buffered saline (1X PBS) (see Reagents and Solutions)

• Calcium sulfate (Drierite, Fisher Scientific, cat. No. 23–116582) or silica desiccant (Silica gel, 2–5 mm, Merck CAS-No:7631–86-9 or equivalent chemical desiccant)

• 1.5-mL sterile microcentrifuge tubes (or equivalent containers for collecting bacteria by centrifugation)

• 0.2-mL 8-tube strips (or equivalent containers for drying cellular reagents)

• Tube racks

• Sterile 100-mm petri dishes

• Plastic 1-mL cuvettes

• 18-gauge needle (optional)

• Parafilm or hot glue gun (optional)

• Sterile spreader or glass beads (for plating bacteria)

• Sterile bacterial culture tubes with caps

• 250-mL sterile conical culture flask

• Pipettes (1000, 200, and 20 μL) and corresponding pipette tips

• 4 °C refrigerator or ice machine

• 42 °C water bath

• 37 °C incubator and shaker (VWR, New Brunswick^™ Innova^R 44/44R, cat No. 75874–524 or equivalent)

• 600 nm wavelength spectrophotometer

• Tabletop microcentrifuge (or equivalent instrument for collecting bacteria by centrifugation)

• 4 oz jars with screw caps (or equivalent containers for drying cellular reagents)

Protocol steps:

Transformation of an appropriate strain of E. coli competent cells with the protein expression plasmid

• 1Place 50 μL of a suspension of competent E. coli in a sterile 1.5-mL microcentrifuge tube (or an equivalent container) and add 1 μL of plasmid DNA.

  Keep competent cells cold at this stage by keeping them on ice or in a 4 °C refrigerator. If starting from a frozen stock of competent cells, thaw them on ice.

  Handle competent cells gently in steps 1–4 and do not subject them to vigorous pipetting or vortexing.

• 2Mix by gently flicking the suspension once or twice and then incubate the tube on ice for 10 to 30 min.
• 3Transfer the tube to a 42 °C water bath and heat-shock the bacteria for 30 sec.
• 4Remove the bacteria from the water bath and place immediately on ice, for 2 min.
• 5Add 250 μL of SOC medium (without any antibiotics) to the bacterial suspension and let the bacteria recover in a 37 °C shaker at 250 rpm for 1 h.
• 6Using a sterile spreader or glass beads, plate 5–10 μL of the transformed bacteria on a selective agar plate containing antibiotic(s). Incubate at 37 °C for 18 to 20 h.

  Use LB agar (or equivalent rich agar medium (Elbing & Brent, 2019)) for this step, and supplement it with the proper plasmid selection antibiotic(s), such as 100 μg/mL of carbenicillin (or ampicillin) or 50 μg/mL of kanamycin.

  In some cases, such as if your protein of interest is toxic and its expression plasmid is leaky, bacterial growth temperature may need to be reduced.

• 7The next day, transfer a single transformant colony into a sterile culture tube containing 3 mL of Superior Broth^™ supplemented with the plasmid selection antibiotic(s). Incubate the culture tube overnight for 18 to 20 h in a 37 °C incubator at 250 rpm.

  Superior Broth^™ may be substituted with alternate rich media, such as LB.

  In some cases, such as if your protein of interest is toxic, bacterial overnight growth temperature may need to be reduced.

Sub-culturing and protein induction

• 8Transfer 250 μL of the overnight bacterial culture into a sterile 250-mL Erlenmeyer flask containing 50 mL of Superior Broth^™ (or equivalent rich media) supplemented with the plasmid selection antibiotic(s).
• 9Incubate this 1:200 bacterial sub-culture in a 37 °C, 250 rpm shaker until its absorbance at 600 nm wavelength (A₆₀₀) is within the range desired for inducing protein expression.

  Measure absorbance of the bacterial culture directly, without dilution, using a spectrophotometer. If a spectrophotometer is not available, McFarland standards may be used to assess culture density (see Alternate Protocol 2).

  For most protein expression plasmids, an A₆₀₀ value of 0.5 to 0.7 is well suited for inducing protein expression. Most 1:200 E. coli sub-cultures usually reach this density within 2 to 2.5 h of aerated growth at 37 °C. Some expression plasmids may give higher protein yields when induced at an A₆₀₀ of ~1.0. We recommend using protein induction conditions reported for previously described plasmids, and for new constructs, empirically determining optimum bacterial culture density for protein induction (see Strategic Planning).

  Adjust bacterial sub-culture conditions, such as sub-culture ratio and growth temperature as necessary, e.g., plasmids encoding toxic proteins may need to be cultured at a lower temperature.

  If the incubator does not have a shaker, incubate cultures in an Erlenmeyer flask with no more than 1/10th volume of broth and shake manually every 15 min. If no incubator is available, a cooler box may be configured into an incubator, as described in detail at https://github.com/FOSH-following-demand/Incubator.

• 10Induce protein expression by adding an appropriate amount of the inducer(s).

  The Lac operator/repressor and the tetracycline promoter/operator systems are two of the most common transcription control systems used to regulate protein expression in E. coli. Transcription from the Lac system is typically induced by adding Isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM, and the tetracycline promoter is usually induced with anhydrotetracycline at a final concentration ranging from 20 to 200 ng/mL. Optimum protein induction parameters may require some trial and adjustments according to the design constraints of the protein and its expression plasmid(s).

  During optimization, protein expression may be verified by methods such as SDS-PAGE analysis of lysed bacteria (Support Protocol 1 and (Gallagher, 2012)) or via functional assays of proteins, using small amounts of lysed bacteria or clarified lysates directly in appropriate reactions (Bhadra, Pothukuchy, et al., 2018).

• 11Continue incubation for 3 to 18 h in a shaking incubator set at 250 rpm and at the desired temperature.

  Protein induction may be done at 37 °C for 3–4 h or at 16 °C to 18 °C for 18–20 h.

  If the incubator does not have a shaker, incubate cultures in an Erlenmeyer flask with no more than 1/10th volume of broth and shake manually every 15 min. If no incubator is available, a cooler box may be configured into an incubator, as described in detail at https://github.com/FOSH-following-demand/Incubator.

Bacteria washing

• 12After completing protein induction, measure absorbance (A₆₀₀) of a 1:10 dilution of the culture.

  Undiluted induced cultures usually reach an A₆₀₀ value >1.0.

  Use this culture density measure as guide for determining the amount of culture to be used for preparing cellular reagents (see Step 13).

• 13Transfer the desired number of 1.2-mL aliquots of the induced culture into 1.5-mL microcentrifuge tubes and pellet the bacteria for 1 min at 9000 × rcf.

  The volume of bacterial culture processed depends on the desired amount of cellular reagents. For most proteins, 5 mL of a culture induced for 3 h and with an A₆₀₀ value of 2.0 will yield cellular reagents for ~500 reactions. Meanwhile, for many proteins induced overnight, 5 mL of the cultures with an A₆₀₀ of 2.0 could yield cellular reagents for ~500 to ~5000 reactions.

  If pelleting bacteria from larger culture volumes, use appropriate centrifugation apparatus and procedures. Refrigeration is not required when centrifuging cultures in a microcentrifuge for 1 min at 9000 × rcf. Longer duration or higher speed of centrifugation might necessitate refrigeration to protect samples from heating.

  If the ambient laboratory temperature during preparation of cellular reagents is significantly greater than 37 °C, bacterial suspensions, especially those expressing thermolabile proteins, may need to be kept on ice or at 4 °C during handling.

• 14Remove the supernatant and wash the bacterial pellet in each tube by resuspending it in 1 mL of cold (4 °C) 1X PBS by gently pipetting up and down.
• 15Centrifuge the tubes at 9000 × rcf for 1 min to pellet the bacteria.
• 16Repeat steps 14 and 15.
• 17Resuspend the bacterial pellets in cold 1X PBS by gently pipetting up and down. Adjust the resuspension volume and pool bacterial pellets as needed so that the final A₆₀₀ value of the suspension ranges between 6.0 and 10.0.

  Use 1:10 dilutions of the bacterial suspensions for measuring the A₆₀₀ values.

Preparation of cellular reagents

• 18Distribute the bacterial suspension into several 8-tube strips of 0.2-mL tubes by aliquoting ~2 × 10⁷ or 2 × 10⁸ washed bacteria per 0.2 mL tube (Figure 2A).
  Prepare a 1:10 dilution of the final suspension of washed bacteria in 1X PBS (from Step 17) before aliquoting 2 × 10⁷ bacteria/tube. Calculate the aliquot volume containing the 2 × 10⁷ or 2 × 10⁸ bacteria by using the final A₆₀₀ value of the washed bacterial suspension and the following approximation: 0.5 A₆₀₀ = 5 × 10⁸ bacteria/mL. For instance, if the final A₆₀₀ value of the washed bacterial suspension in Step 17 is 6.5, 31 μL of the bacterial suspension will contain 2 × 10⁸ bacteria, as determined by using the equation 1:

  $$\frac{1000\mu \text{L}}{{65\times 10}^8\mathit{\text{ bacteria}}}=\frac{\mathit{\text{aliquot volume }}\left(\mathit{\text{in }}\mu \text{L}\right)}{{2\times 10}^8\mathit{\text{ bacteria}}}.$$
  Consequently, the number of 8-tube strips needed to distribute the entire volume of bacterial suspension can be determined by solving equation 2:

  $$\left(\frac{\mathit{\text{Volume of bacterial suspension }}\left(\mathit{\text{in }}\mu L\right)}{\mathit{\text{aliquot volume }}\left(\mathit{\text{in }}\mu L;\mathit{\text{ from equation }}1\right)}\right)/8$$

  Optional: If it is desired that cellular reagent preparations do not contain live bacteria, for instance, to abide shipping or distribution regulations, you can inactivate bacteria while maintaining enzymatic function in cellular reagent preparations, by incubating the bacterial aliquots at 60 °C for 10 min.

  For thermolabile proteins that cannot tolerate 60 °C, however, one of the ways in which cellular reagent preparations lacking live bacteria may be prepared, is by lyophilization (Bhadra, Pothukuchy, et al., 2018) (see Alternate Protocol 1).

• 19Fill an airtight container that can accommodate several 8-tube strips of 0.2-mL tubes (such as a screw cap plastic jar or Tupperware with a lid) halfway with a chemical desiccant such as calcium sulfate (commercially known as Drierite^™, which is available in both dryness indicating and non-indicating forms).

  Do not leave the desiccant exposed to air for a long time. Some of the commonly used desiccants, such as indicating Drierite^™ (Fisher Scientific, cat. No. 23–116582), are impregnated with cobalt chloride that acts as a color indicator of dryness, by turning from blue to pink upon absorption of moisture.

• 20Place the uncapped tubes containing the bacterial aliquots upright inside the desiccant-filled container and then tightly close the container lid.

  Do not overfill a desiccant container with bacterial aliquot. Typically, a container filled with 250 g of calcium sulfate desiccant should hold at most four 8-tube strips of 0.2-mL tubes, each containing up to 30–35 μL liquid/tube (Figure 2B).

  Note the volume of bacterial suspension/0.2 mL tube that was aliquoted in Step 18 on the container lid to aid subsequent rehydration for use in subsequent applications (see Basic Protocols 2 and 3).

• 21Place the container in a 37 °C incubator for 18–24 h to dry the bacterial suspensions.

  Keep the desiccant container upright and do not get any desiccant inside the bacterial tubes (Figure 2C). If creating a small opening in the tubes makes handling easier, cap the bacteria-filled tubes and create a small hole for evaporation by gently poking through the cap with an 18-gauge needle. Then place these tubes in the desiccant container and tightly close the container lid. This hole may be sealed after the bacteria are dry by either wrapping the tubes in parafilm or by placing a drop of hot glue over the hole using a hot glue gun.

  Visually inspect the bacterial suspensions for dryness. Continue incubation at 37 °C till no liquid can be seen in the tubes and the bacteria appear as a yellowish powdery coating on the tubes (Figure 2D).

  37 °C is recommended as the drying temperature because of the convenience of maintaining this temperature using the same equipment that was used to grow the protein-expressing bacterial culture. If desired, bacterial suspensions may be dried at room temperature (25 °C), however, longer incubation times are usually necessary.

• 22Close the tube caps and place them back inside the airtight desiccant container.
• 23Store the cellular reagent containers in a dark cabinet at room temperature or inside a 4 °C refrigerator until use.

Figure 2. Images depicting cellular reagents in various stages of preparation and use.

(A) Aliquots of washed protein-expressing bacteria dispensed in 8-strip 0.2-mL tubes. (B) Tubes containing bacterial suspensions placed with their caps open inside a plastic container filled halfway with desiccant pellets. (C) Tightly capped plastic container with open tubes of bacterial suspension and desiccant pellets ready for incubation at 37 °C. (D) Dry cellular reagents after overnight incubation at 37 °C. I Cellular reagents rehydrated in 30 μL of water.

ALTERNATE PROTOCOL 1: PREPARATION OF LYOPHILIZED CELLULAR REAGENTS

Cellular reagents prepared using Basic Protocol 1 usually contain some viable bacteria (Bhadra, Nguyen, et al., 2021). Although over time, and during storage, bacterial viability declines, it may be necessary in some cases, such as for off-site distribution, to ensure absence of live bacteria in preparations of cellular reagents. This may be readily achieved by heating the bacteria at 60 °C for 10 min prior to drying them at 37 °C (see Basic Protocol 1). However, heat-killing may be unsuitable for thermolabile proteins. In such cases, lyophilization in the absence of excipients such as trehalose may be used to prepare cellular reagents lacking viable E. coli (Bhadra, Pothukuchy, et al., 2018).

This protocol describes how to prepare cellular reagents by freeze-drying protein expressing bacteria using a lyophilizer. The initial steps of bacterial transformation, induction of protein expression, and distribution of washed protein-expressing bacteria into 0.2-mL tubes, as well as methods for long-term storage of the lyophilized cellular reagents, are the same as those described in Basic Protocol 1. Usage methods and applications (Basic Protocols 2 and 3) and performance of lyophilized cellular reagents are also similar to those of cellular reagents prepared using Basic Protocol 1.

Additional Materials to those in Basic Protocol 1:

• Dry ice or −80 °C freezer

• Benchtop lyophilizer (VirTis Benchtop Pro lyophilizer or equivalent)

• Cold PCR tube rack (optional)

Protocol steps:

1. Prepare and aliquot bacteria expressing the protein of interest by following steps 1–18 of Basic Protocol 1.

   When aliquoting washed bacteria into 8-tube strips of 0.2-mL tubes in Step 18, place the tube strips on a cold tube rack or on ice. Distribute volumes containing enough bacteria to perform at least 10 reactions of the molecular biology procedure in which the reagents are intended to be used, into each 0.2-mL tube. Very small aliquots per tube may be difficult to handle since they might partially thaw in the time it takes to load them in the lyophilizer. This, in turn, may lead to reduced activity of the resulting cellular reagents.

2. Cap the 0.2-mL tubes containing the aliquots of protein-expressing bacteria and make a single hole in each cap using an 18-gauge needle (Figure 3).

3. Cover the rack of tubes with a piece of aluminum foil (or an equivalent cover) and freeze the bacterial suspensions for at least one hour by storing in a −80 °C freezer or dry ice.

   Bacteria may be stored frozen overnight prior to lyophilization.

4. Prepare the lyophilizer for use according to the manufacturer’s instructions.

   For the VirTis Benchtop Pro lyophilizer, use the automatic condenser and vacuum pump settings to maintain the instrument at 197 to 215 mTorr and −108 °C.

5. When the lyophilizer has reached the appropriate vacuum and temperature measurements and is ready to freeze-dry samples, place the tubes of frozen bacteria in the lyophilizer and immediately start the freeze-drying process.

6. Lyophilize the bacterial suspensions for ~3 h.

   Freeze-dried cellular reagents have a white powdery appearance (Figure 3).

7. Release the lyophilizer vacuum according to the manufacturer’s instructions and collect the tubes containing dried cellular reagents.

8. Seal the holes in the tube caps with parafilm or a dab of hot glue applied using a hot glue gun.

9. Place the cellular reagents inside a desiccant-filled container and store in a dark cabinet or in a 4 °C refrigerator until use.

Figure 3. Images depicting some stages in the preparation of lyophilized cellular reagents.

(A) Use of an 18-gauge needle to poke a hole in the cap of 0.2-mL tubes containing aliquots of the bacterial suspension. (B) Lyophilized cellular reagents.

ALTERNATE PROTOCOL 2: EVALUATION OF BACTERIAL CULTURE GROWTH VIA COMPARISON TO McFARLAND TURBIDITY STANDARDS

The following protocol describes an approach that should allow preparation of cellular reagents in the absence of a spectrophotometer. As described in Basic Protocol 1, a spectrophotometer is used to track bacterial growth by measuring absorbance of the culture at 600 nm wavelength. If a spectrophotometer is not available, McFarland turbidity standards may be used to assess the density of bacterial cultures (Baldwin et al., 2012; McFarland, 1907; Voigt & Voigt, 2011), by performing visual comparisons of the opacity of the cultures and the standards. These turbidity standards are chemical solutions of barium chloride and sulfuric acid, which react to form fine precipitates of barium sulfate. When shaken well, these precipitate suspensions visually mimic bacterial suspensions in opacity.

Materials:

• 1% (wt/vol) solution of anhydrous barium chloride (BaCl₂, 0.048 M, Sigma, cat. No. 342920)

• 1% (vol/vol) solution of sulfuric acid (H₂SO₄, 0.18 M, Sigma, cat. No. 258105)

• Paper with pattern of alternating black and white lines (see Figure 4). Alternating pattern of ~0.25 inches thick black and white lines may be created using drawing tools such as those available in Microsoft PowerPoint or equivalent, and printed on white paper. Alternatively, a plastic laminated Wickerham card with a thick pattern of black and white lines may be purchased from commercial sources, such as VWR, cat. No. 76288–330.

• Borosilicate glass tubes (VWR, cat No. 490015–324, or equivalent)

• Tube rack that allows tubes to be viewed from the side

Figure 4. Measurement of bacterial culture density using McFarland turbidity standards.

(A) McFarland turbidity standards representing bacterial cultures with A₆₀₀ values in the range of 0.1 to 1.0. (B) Comparison of a bacterial culture with a A₆₀₀ value of 0.38 with McFarland standards representing A₆₀₀ values ranging from 0.2 to 0.6. (C) Pattern of horizontal black and white lines for comparing turbidity.

Protocol steps:

1. Prepare 11 standard solutions by mixing a 1% (wt/vol) solution of barium chloride and a 1% (vol/vol) solution of sulfuric acid in the ratios shown in Table 1.

2. Aliquot each standard into clear tubes of the same size and shape such that the tubes are filled to a height of about 2.5 cm.

   Vigorously vortex or shake the standards before use, and visually ensure that they are uniformly turbid.

   Cap the tubes tightly and keep the standards in the dark when not in use for at most 12 weeks.

3. Place the tubes in a rack that allows the tubes to be viewed from the side.

4. Place the paper with the pattern of alternating black and white lines behind the standards (Figure 4).

5. Aliquot the bacterial culture to be measured to a height of 2.5 cm into the same type of tube that was used to prepare the standards.

6. Place the tube containing the culture next to the standards and make a side-by-side comparison of how much the black and white pattern is obscured by the solutions in the tubes.

7. Use the standard that obscures the pattern to the same extent as the bacterial culture as the best estimation of sample turbidity.

Table 1.

Preparation of McFarland turbidity standards and corresponding OD₆₀₀ value.

1% (wt/vol) BaCl2 (μL)a	1% (vol/vol) H2SO4 (μL)	%vol/vol of 1% BaCl2 in total volume	Corresponding OD600
0	3,000	0	0
10.89	2,989	0.363	0.1
21.78	2,978	0.726	0.2
32.67	2,967	1.089	0.3
43.56	2,956	1.452	0.4
54.45	2,945	1.815	0.5
65.34	2,934	2.178	0.6
76.23	2,923	2.541	0.7
87.12	2,912	2.904	0.8
98.01	2,901	3.267	0.9
108.9	2,891	3.63	1

^aThe indicated μL quantities are for the preparation of 3 mL of standards; the volumes can be scaled up or down accordingly.

SUPPORT PROTOCOL 1: SDS-PAGE FOR PROTEIN EXPRESSION ANALYSIS OF CELLULAR REAGENTS

The following protocol describes the use of SDS-PAGE to analyze the expression level of the desired recombinant protein in bacteria, for use in Basic Protocol 1. SDS-PAGE may be used both to guide optimization of protein expression conditions for preparation of cellular reagents and to confirm protein expression during preparation of cellular reagents.

Materials:

• Parallel bacterial cultures transformed with the plasmid of interest and grown in the presence (induced) or absence (uninduced) of the inducer of protein expression

• Untransformed bacterial culture grown under the same conditions and duration as the transformed bacterial cultures

• 1X Phosphate buffered saline (1X PBS) (see Reagents and Solutions)

• SDS-PAGE sample buffer (see Reagents and Solutions)

• 12% Bis-Tris SDS-PAGE precast gel and running buffer (Invitrogen, cat. No. NP0341PK2, or equivalent). SDS-PAGE gels and running buffers may be prepared in-house using protocols described in (Gallagher, 2012).

• Multicolor broad range protein ladder (Thermo Scientific, cat. No. 26623, or equivalent)

• Gel code Blue (Thermo Scientific, cat. No. 24590, or equivalent)

• Dry heat block (VWR, cat No. 75838–286, or equivalent)

• Tabletop microcentrifuge (or equivalent instrument for collecting bacteria by centrifugation)

Protocol steps:

1. Place aliquots of the untransformed (not expressing protein of interest), uninduced (expressing none to low levels of the protein of interest), and induced (expressing protein of interest) bacterial cultures in 1.5-mL microcentrifuge tubes.

   Bacteria may be harvested at the same time from two duplicate (parallel) cultures of which only one was treated with inducers of protein expression (Basic Protocol 1, steps 8–11).

   If the expression system being used is expected to be leaky and produces some protein even in the absence of any inducers, bacteria that have not been transformed with the expression plasmid may also be analyzed, for clarity of results. If, alternatively, bacteria are being harvested from the same culture before and after induction, ensure that equivalent bacterial amounts, as extrapolated from the A₆₀₀ values, are used for comparison of protein expression.

   The number of bacteria used for analysis can vary depending on the protein expression level and detection method. In general, 2 × 10⁸ to 10 × 10⁸ bacteria (~1 mL of culture at 0.2 to 1.0 A₆₀₀) may be sufficient for SDS-PAGE analysis.

2. Pellet bacteria in a microcentrifuge at 9000 × rcf for 1 min, discard the cell culture supernatant, and wash the pellet twice with 1X PBS.

   Bacterial pellets may be stored at −20 °C to −80 °C for a few days.

3. Resuspend the bacterial pellets in 25–50 μL of SDS-PAGE sample buffer.

4. Lyse the samples by incubating at 95 °C for 5–10 min.

5. Centrifuge the samples at 13,000 × g for 2 min and transfer the supernatants to new microcentrifuge tubes.

6. Run 10–20 μL of the samples and a suitable protein ladder on a precast 12% Bis-Tris SDS-PAGE gel according to the manufacturer’s suggestions, or on an in-house prepared gel, as detailed in (Gallagher, 2012).

7. Stain the gel with Gel code blue or equivalent Coomassie stain according to the manufacturer’s instructions.

   See example in Understanding Results.

BASIC PROTOCOL 2: USING TAQ DNA POLYMERASE CELLULAR REAGENTS FOR PCR

As a specific example of opportunities for cellular reagent applications, this protocol describes the execution of PCR, one of the most used nucleic acid amplification techniques, using cellular reagents. PCR amplifies DNA templates using two primers, a thermostable DNA polymerase, and multiple cycles of stepwise changes in temperature that lead to iterative DNA denaturation, primer annealing, and polymerase-mediated primer extension. The accumulating PCR amplicons may be analyzed in real-time using DNA binding dyes or nucleic acid probes, for applications in diagnostics or detection. PCR products may also be visualized at end point, for instance, by using agarose gel electrophoresis, and may then be used in downstream reactions, such as DNA assembly (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018). In this protocol, Taq DNA polymerase cellular reagents produced following Basic Protocol 1 are rehydrated and used instead of pure Taq DNA polymerase for performing a real-time PCR assay designed to amplify a region of Chlamydia trachomatis 16S rRNA gene (Table 2). This assay may be employed by users as a positive control to assess their preparations of Taq DNA polymerase cellular reagents. This protocol should serve as a general guideline for the application of Taq DNA polymerase cellular reagents in other PCR assays.

Table 2.

PCR primer sequences, EvaGreen qPCR recipe, and qPCR cycling conditions for the detection of Chlamydia trachomatis 16S rDNA sequence.

Primer name	Sequence
CT16S.Fwd	5’-TAGTGGCGGAAGGGTTAG-3’
CT16S.Rev	5’-CGTCATAGCCTTGGTAGG-3’
C. trachomatis 16S rDNA PCR amplicon sequence	5’-TAGTGGCGGAAGGGTTAGTAATGCATAGATAATTTGTCCTTAACTTGGGAATAACGGTTGGAAACGGCCGCTAATACCGAATGTGGCGATATTTGGGCATCCGAGTAACGTTAAAGAAGGGGATCTTAGGACCTTTCGGTTAAGGGAGAGTCTATGTGATATCAGCTAGTTGGTGGGGTAAAGGCCTACCAAGGCTATGACG-3’
EvaGreen qPCR recipe
Component	Volume (μL) per 25 μL reaction
10X ThermoPol buffer	2.5
Template DNAa	3
10 μM CT16S.Fwd	1.25
10 μM CT16S.Rev	1.25
4 mM dNTP mix	2.5
20X EvaGreen	1.25
2 × 107 Taq DNA polymerase dried bacteria / 3 μL	3
Water	Up to 25 μL
	PCR cycling conditions
Step 1: 95 °C	10 min
Step 2: 95 °C	10 sec
Step 3: 55 °C	15 sec
Step 4: 72 °C	30 sec

^aA 10-fold dilution series of plasmid or gBlock^™ (IDT) templates starting from 2 × 10⁶ copies/μL to 2 × 10² copies/μL may be used. No template controls should receive an equal amount of water or non-specific templates.

Set the qPCR machine to perform 45 cycles of amplification (Steps 2 to 4, with EvaGreen fluorescence measured in Step 4 of each cycle) followed by melt curve analysis of the PCR amplicons.

Materials:

• Taq DNA polymerase dried bacteria obtained following Basic Protocol 1 using BL21 bacteria transformed with plasmid pATetO 6XHis-Taq DNA Pol (Addgene #178782)

• Amplification buffer (10X ThermoPol or equivalent buffers; see Reagents and Solutions)

• Deoxyribonucleotides (dNTP) (NEB, cat. No. N0446S, or equivalent)

• Nuclease-free water

• Nucleic acid samples to test (water or TE 10:0.1 buffer containing plasmids or double stranded DNA fragments such as gBlocks^™ or eBlocks^™ from Integrated DNA Technologies (IDT), bearing the C. trachomatis 16S rDNA amplicon sequence noted in Table 2, may be used)

• Oligonucleotide primers (see example sequences in Table 2)

• EvaGreen (Biotium, cat. No. 31000) or equivalent DNA intercalating dye (Mackay & Landt, 2007)

• Barrier tips for pipettes

• Real-time PCR tubes or plates

• Real-time PCR machine (LightCycler96, Roche, or equivalent)

Protocol steps:

1. Assemble a PCR master mix on ice or at 4 °C by combining all reaction components (see Table 2 for qPCR recipe), except the template and the cellular reagents. Prepare enough master mix to analyze five different template copy numbers and one no template control, all in triplicate.

   Maintain the master mix at 4 °C or on ice at all times prior to starting the amplification process.

   For use as a positive control for validating preparations of Taq DNA polymerase cellular reagents, qPCR reaction composition and cycling conditions for amplifying a sequence from Chlamydia trachomatis 16S rDNA gene are provided in Table 2. The C. trachomatis PCR amplicon sequence is also noted in Table 2. Synthetic double stranded DNA fragments (eBlocks^™ or gBlocks^™, IDT) bearing this sequence may be used as templates either directly or after cloning into a plasmid. If desired, this PCR assay can also be used to amplify this 16S rDNA sequence from C. trachomatis genomic DNA (ATCC, cat. No. VR-885D).

   Ensure that all individual reagents, especially those that are thawed from −20 °C, such as the reaction buffer and dNTPs, are uniformly mixed before use.

2. Rehydrate the desired number of Taq DNA polymerase cellular reagent tubes, each containing about 2 × 10⁸ dried bacteria (see Basic Protocol 1), with 30 μL of water by pipetting gently up and down.

   For triplicate C. trachomatis 16S qPCR analysis of five template amounts and one no template control, 3.6 × 10⁸ dried bacteria will be required. Therefore, rehydration of two tubes of Taq DNA polymerase cellular reagents with 2 × 10⁸ dried bacteria in each tube should be sufficient to perform the assay.

   The volume of water used to rehydrate the cellular reagents may need to be adjusted to match the starting volume of the bacterial suspension that was added to the tube during preparation of the cellular reagents (and noted on the cellular reagent container; see Basic Protocol 1).

   Cellular reagents should be rehydrated immediately before use. Do not store rehydrated cellular reagents for later use.

3. Transfer the desired volume of rehydrated cellular reagents into the PCR master mix.

   In most protocols, cellular reagents are used at 2 × 10⁶ to 2 × 10⁷ dried bacteria/reaction. As described in Basic Protocol 1, this is not an exact number based on bacterial plate counts but rather an extrapolated number derived from the final A₆₀₀ value of the washed bacterial suspension used to prepare the cellular reagent and the approximation 0.5 A₆₀₀ = 5 × 10⁸ bacteria/mL. The amount of cellular reagents that is required in a reaction usually depends on the strength and duration of protein induction performed during preparation of the cellular reagent. The amount used is also contingent on the type of reaction. For instance, a 20-μL Gibson assembly reaction usually contains 2 × 10>⁷ Taq DNA ligase cellular reagents, but only 1 × 10⁵ and 1 × 10⁶ of similarly prepared T5 exonuclease and Taq DNA polymerase cellular reagents, respectively. Since both too many and too few cellular reagents in a reaction can be detrimental, users should experimentally determine the optimal application amount for their cellular reagent preparations.

   About 2 × 10⁷ dried bacteria of Taq DNA polymerase cellular reagents prepared after a 3 h induction of protein expression from a tetracycline promoter perform well for most endpoint and real-time PCR assays. Therefore, use 1/10^th of the volume of the rehydrated 2 × 10⁸ dried bacteria (3 μL from a 30 μL rehydration volume) for each PCR in order to achieve 2 × 10⁷ dried bacteria/reaction.

   Once the cellular reagents are added, the PCR reactions should be processed for amplification and readout as quickly as possible. Do not store fully assembled assays for long durations.

4. Uniformly mix the PCR master mix by gentle vortexing or pipetting up and down, and then transfer the desired volume per reaction into individual thin-walled 0.2 mL-reaction tubes or 96-well plates.

   Typically, PCR is performed in a total volume of 25 μL to 50 μL that includes 2 to 5 μL of the template.

5. Prepare the ‘no template’ negative controls for assessing amplification specificity by adding the required volume of a non-specific template or water to the proper PCR reactions and then cap or cover these reactions.

6. Prepare the DNA templates as desired and, if necessary, dilute them in TE 10:0.1 buffer or water.

   Template copies used may be varied from single digits to several hundreds of thousands per reaction depending on the detection limit of the assay, the readout method (for instance, endpoint ethidium bromide agarose gel electrophoresis usually needs more template input as compared to real-time PCR), and the number of PCR amplification cycles to be used.

   For the C. trachomatis 16S qPCR assay, prepare five different concentrations of double stranded DNA templates, containing 2 × 10⁶, 2 × 10⁵, 2 × 10⁴, 2 × 10³, or 2 × 10² copies/μL.

7. Add the required amount of template to the proper PCR reactions and incubate them immediately in a thermocycler programmed to incubate the reactions through the desired temperature cycling conditions.

   To prepare a standard curve for the C. trachomatis 16S qPCR assay, add the following number of template copies/reaction in triplicate PCR assays: 6 × 10⁶, 6 × 10⁵, 6 × 10⁴, 6 × 10³, and 6 × 10², and analyze the PCR assays in a real-time PCR machine.

   Taq DNA polymerase cellular reagent-based PCR assays may be analyzed at endpoint, using agarose gel electrophoresis, or measured in real-time, using DNA binding dyes such as EvaGreen (see Figure 5) or TaqMan probes.

Figure 5. Using Taq DNA polymerase cellular reagents for PCR.

(A) Schematic depicting the applications of Taq DNA polymerase cellular reagents in endpoint and real-time quantitative PCR. (B and C) Real-time quantitative PCR assays to assess batch-to-batch performance of Taq DNA polymerase cellular reagents. A Chlamydia trachomatis 16S rDNA sequence was PCR-amplified in three technical replicates using three different batches of Taq DNA polymerase cellular reagents. Amplification curves generated by measuring real-time increase in EvaGreen fluorescence upon amplification of 6 × 10⁶ (blue traces), 6 × 10⁵ (black traces), 6 × 10⁴ (red traces), 6 × 10³ (pink traces), 6 × 10² (light blue traces), and 0 (gray traces) template starting copies per reaction by the three cellular reagent batches are depicted in panel B. Melting peaks of the resulting PCR amplicons, generated using the ‘Tm calling’ analysis protocol in the LightCycler96 software are depicted in panel C.

BASIC PROTOCOL 3: USING Br512 DNA POLYMERASE CELLULAR REAGENTS FOR LAMP

LAMP is an isothermal amplification method that uses four to six primers and a strand-displacing DNA polymerase (such as the B. stearothermophilus DNA polymerase) to amplify DNA and RNA templates (Nagamine, Hase, & Notomi, 2002; Notomi et al., 2000). The basic mechanism of LAMP results in the production of continuously elongating concatemers with single-stranded loop sequences at both ends that form a ladder-like pattern in agarose gels (Figure 6) (Notomi et al., 2000). LAMP assays can also be analyzed in real-time or at endpoint without opening the reaction tubes (Mori, Kitao, Tomita, & Notomi, 2004; Notomi et al., 2000). Non-specific indicators of amplification, such as increases in turbidity due to Mg²⁺ precipitation, changes in color of pH sensitive dyes, and increases in fluorescence from DNA intercalating dyes, are commonly used for such readout (Tanner, Zhang, & Evans, 2015; Tomita, Mori, Kanda, & Notomi, 2008). However, given the propensity for spurious amplification in continuous amplification methods such as LAMP, the preferred method for observing LAMP amplicons is by using oligonucleotide strand displacement (OSD) hemiduplex DNA probes (Jiang et al., 2015) that, similar to TaqMan probes for PCR, suppress noise and produce a sequence-specific signal. In LAMP-OSD, the long strand of the hemiduplex DNA is labeled with a fluorophore and the short strand is labeled with a quencher (Jiang et al., 2015); the short single-stranded region at one end of the OSD long strand acts as a toehold that initiates strand exchange hybridization with the single-stranded LAMP amplicon loop, which, in turn, leads to displacement of the short OSD strand and a resulting fluorescent signal (Figure 6) that can be measured in real-time or observed at endpoint (Bhadra, Riedel, Lakhotia, Tran, & Ellington, 2021; Bhadra, Riedel, et al., 2018; Bhadra, Saldana, Han, Hughes, & Ellington, 2018; Jiang et al., 2018; Jiang, Stacy, Whiteley, Ellington, & Bhadra, 2017). Herein we describe a LAMP-OSD assay procedure using cellular reagents expressing a superior, engineered strand-displacing DNA polymerase, Br512 Mut235 (Paik et al., 2021). In this protocol, Br512 Mut235 cellular reagents, prepared according to Basic Protocol 1, are rehydrated and used to perform two different LAMP-OSD assays: (1) a multiplex LAMP-OSD assay for detection of SARS-CoV-2 virions and (2) a LAMP-OSD assay for detection of the human glyceraldehyde-3-phosphate dehydrogenase (gapd) gene. Each type of LAMP-OSD assay will be set up as an assay validation test comprising several positive control reactions containing known amounts of specific templates (SARS-CoV-2 virions and synthetic gapd DNA, respectively) and ‘no-template’ negative control reactions lacking specific amplification targets. The reaction recipes may be used for SARS-CoV-2 diagnostics, where the gapd LAMP-OSD assay would serve as an amplification control for human samples, such as heated saliva. Annotations within the protocol provide additional guidance on diagnostic applications. Both protocols may also be used as assessment tools for verifying new preparations of Br512 Mut235 cellular reagents. The steps and notes detailed in this protocol should also serve as general guidelines for the application of Br512 Mut235 cellular reagents in LAMP-OSD assays for other DNA and RNA targets of interest.

Figure 6. Schematics depicting the LAMP-OSD mechanism.

The left panel depicts the formation of concatemeric LAMP amplicons from target nucleic acids using strand displacing DNA polymerase (denoted as Bst) and four LAMP primers (denoted as FIP and BIP for the inner primers and F3 and B3 for the outer primers). The inner primer binding regions in the target are colored-coded and labeled as F1 (red), F2 (black), B1 (blue), and B2 (gray). The corresponding complementary regions in the antisense strand are denoted as F1’ (red striped), F2’ (black striped), B1’ (blue striped), and B2’ (gray striped). The agarose gel image depicts the typical ladder-like pattern of specific and non-specific LAMP amplicons. 3’-ends of DNA are denoted by arrowheads. The right panel depicts formation of hemiduplex OSD probe by annealing the OSD long and short strands labeled with a fluorescein (FAM) and quencher (Q), respectively. Toehold-mediated strand displacement hybridization of the OSD long strand starts by hybridization of its single-stranded toehold (green) to the complementary region in the single-stranded LAMP amplicon loop (green striped). Subsequent branch migration leads to hybridization of the long OSD strand to the LAMP loop and the ensuing separation of the short OSD strand results in fluorescence signal. Image at the bottom (taken with a cell phone) depicts an example of visual yes/no readout of endpoint OSD fluorescence in a LAMP-OSD assay.

Materials:

• Br512 Mut235 DNA polymerase dried bacteria obtained following Basic Protocol 1 using BL21(DE3) bacteria transformed with plasmid pKAR2-Br512g2.2 (Addgene # 179278)

• Amplification buffer (10X G6 or equivalent buffers; see Reagents and Solutions)

• Deoxyribonucleotides (dNTP) (NEB, cat. No. N0446S, or equivalent)

• Nuclease-free water

• Oligonucleotide primers (see example sequences and preparation instructions in Tables 3 and 4)

• OSD probes (see Reagents and Solutions and Tables 3 and 4 for sequences and preparation instructions)

• Nucleic acid samples to test: inactivated SARS-CoV-2 virions (BEI Resources, cat. No. NR-52287, or equivalent) and plasmids or synthetic double stranded DNA (such as gBlocks^™ or eBlocks^™, IDT) bearing the gapd LAMP amplicon sequence noted in Table 4. Nucleic acid samples may range from purified nucleic acids in water or TE 10:0.1 buffer to more complex materials, such as virions in culture supernatants or TE 10:0.1 buffer, or even crude samples, such as heated human saliva containing virions and bacteria.

• 5M betaine solution (Sigma, cat. No. B2300, or equivalent)

• Barrier tips for pipettes

• Thin-walled 0.2-mL tubes with no auto-fluorescence in the fluorescein range as verified by visual inspection of empty tubes in a blue light transilluminator, gel doc, or DIY fluorescence visualization device (see Support Protocol 2) (VWR, cat. No. 10011–816, or equivalent)

• Thermocycler or dry heat block (VWR, cat. No. 75838–286, or equivalent)

• Blue light transilluminator, gel doc, or DIY fluorescence visualization instrument (see Support Protocol 2)

Table 3.

Multiplex LAMP-OSD assay primers, OSD probes, and reaction recipe for detection of SARS-CoV-2.

Primer/OSD probe name	Sequence
NB-F3	5’-ACCGAAGAGCTACCAGACG-3’
NB-B3	5’-TGCAGCATTGTTAGCAGGAT-3’
NB-FIP	5’-TCTGGCCCAGTTCCTAGGTAGTTCGTGGTGGTGACGGTAA-3’
NB-BIP	5’-AGACGGCATCATATGGGTTGCACGGGTGCCAATGTGATCT-3’
NB-LB	5’-ACTGAGGGAGCCTTGAATACA-3’
NB-OSD-F	/56-FAM/CCG AAT GAA AGA TCT CAG TCC AAG ATG GTA TTT CT/3InvdT/
NB-OSD-Q	5’-TC TTG GAC TGA GAT CTT TCA TTC GG /3IABkFQ/
Lamb-F3	5’-TCCAGATGAGGATGAAGAAGA-3’
Lamb-B3	5’-AGTCTGAACAACTGGTGTAAG-3’
Lamb-FIP	5’-AGAGCAGCAGAAGTGGCACAGGTGATTGTGAAGAAGAAGAG-3’
Lamb-BIP	5’-TCAACCTGAAGAAGAGCAAGAACTGATTGTCCTCACTGCC-3’
Lamb-LF	5’-CTCATATTGAGTTGATGGCTCA-3’
Lamb-LB	5’-ACAAACTGTTGGTCAACAAGAC-3’
Lamb-OSD-F	5’-GTA TGG TAC TGA AGA TGA TTA CCA AGG TAA ACC TTT GGA ATT TGG AC/36-FAM/
Lamb-OSD-Q	/5IABkFQ/GT CCA AAT TCC AAA GGT TTA CCT TGG TAA TCA TCT C/3InvdT/
Multiplex LAMP-OSD recipe
Component	Volume (μL) per 25 μL reaction
RNA or virion templatea	3
10 mM dNTP mix	3.5
NB LAMP primersb	1.5
Lamb LAMP primersc	1.5
10X G6D buffer	2.5
5M Betaine	2
NB OSD probesd	2.5
Lamb OSD probese	2.5
2 × 107 Br512 Mut235 dried bacteria / 3 μL	3
water	Up to 25 μL

^aUse ≥500 copies of RNA or virion templates per reaction.

^bNB LAMP primers (Zhang et al., 2020) comprise a mix of 25 μM each of NB-FIP and NB-BIP, 10 μM of NB-LB loop primer, and 5 μM each of NB-F3 and NB-B3 primers in TE 10:0.1 buffer.

^cLamb LAMP primers (Lamb, Bartolone, Ward, & Chancellor, 2020) comprise a mix of 20 μM each of Lamb-FIP and Lamb-BIP, 10 μM each of Lamb-LB and Lamb-LF loop primers, and 5 μM each of Lamb-F3 and Lamb-B3 primers in TE 10:0.1 buffer.

^dNB OSD probes are composed of 1 μM of the fluorescein-labeled strand (NB-OSD-F) annealed with 5 μM of the quencher-labeled strand (NB-OSD-Q).

^eLamb OSD probes are composed of 1 μM of the fluorescein-labeled strand (Lamb-OSD-F) annealed with 3 μM of the quencher-labeled strand (Lamb-OSD-Q).

Oligonucleotide primers and probes may be purchased from IDT. /3IABkFQ/ and /5IABkFQ/ are the notations used by IDT to represent Iowa Black^®FQ dark quencher at the 3’-end or 5’-end, respectively, of an oligonucleotide. /3InvdT/ is the notation used by IDT to represent a 3’-end inverted dT that prevents extension by DNA polymerases. /36-FAM/ and /56-FAM/ are the notations used by IDT to represent a single isomer derivative of fluorescein attached to the 3’-end or 5’-end, respectively, of an oligonucleotide.

Table 4.

LAMP-OSD assay primers, OSD probes, and reaction recipe for detection of human glyceraldehyde 3-phosphate-dehydrogenase gene.

Primer/OSD probe name	Sequence
hGAPD.F3	5’-GCCACCCAGAAGACTGTG-3’
hGAPD.B3	5’-TGGCAGGTTTTTCTAGACGG-3’
hGAPD.FIP	5’-CGCCAGTAGAGGCAGGGATGAGGGAAACTGTGGCGTGAT-3’
hGAPD.BIP	5’-GGTCATCCCTGAGCTGAACGGTCAGGTCCACCACTGACAC-3’
hGAPD.LR	5’-TGTTCTGGAGAGCCCCGCGGCC-3’
hGAPD.OSD.F	/56-FAM/CTCACTGGCATGGCCTTCCGTGTCCCCACTGCCAAC/3InvdT/
hGAPD.OSD.Q	5’-GGACACGGAAGGCCATGCCAGTGAG/3IABkFQ/
GAPD LAMP amplicon sequence	5’-GCCACCCAGAAGACTGTGGATGGCCCCTCCGGGAAACTGTGGCGTGATGGCCGCGGGGCTCTCCAGAACATCATCCCTGCCTCTACTGGCGCTGCCAAGGCTGTGGGCAAGGTCATCCCTGAGCTGAACGGGAAGCTCACTGGCATGGCCTTCCGTGTCCCCACTGCCAACGTGTCAGTGGTGGACCTGACCTGCCGTCTAGAAAAACCTGCCA-3’
LAMP-OSD recipe
Component	Volume (μL) per 25 μL reaction
DNA templatea	5
10X G6B buffer	2.5
4 mM dNTP mix	2.5
LAMP primer mixb	2
OSD probesc	2.5
5 M Betaine	5
2 × 107 Br512 Mut235 dried bacteria / 3 μL	3
Water	Up to 25 μL

^aUse ≥600 copies of DNA templates per reaction.

^bLAMP primer mix comprises 20 μM each of hGAPD.FIP and hGAPD.BIP, 10 μM of hGAPD.LR loop primer, and 5 μM each of hGAPD.F3 and hGAPD.B3 primers in TE 10:0.1 buffer.

^cOSD probes are composed of 1 μM of the fluorescein-labeled strand (hGAPD.OSD.F) annealed with 2 μM of the quencher-labeled strand (hGAPD.OSD.Q).

Oligonucleotide primers and probes and double stranded DNA gBlock^™ or eBlock^™ templates may be purchased from IDT. /3IABkFQ/ is the notation used by IDT to represent Iowa Black^®FQ dark quencher at the 3’-end of an oligonucleotide. /3InvdT/ is the notation used by IDT to represent a 3’-end inverted dT that prevents extension by DNA polymerases. /56-FAM/ is the notation used by IDT to represent a single isomer derivative of fluorescein attached to the 5’-end of an oligonucleotide.

Protocol steps:

1. Assemble the SARS-CoV-2 and gapd LAMP-OSD reaction master mixes noted in Tables 3 and 4, respectively, on ice or at 4 °C by combining all reaction components except the templates and the cellular reagents.

   Maintain the LAMP-OSD reaction mixes at 4 °C or on ice at all times prior to starting amplification at 65 °C.

   Prepare both master mixes in sufficient amounts to set up four positive control reactions to which different amounts of specific nucleic acid targets (SARS-CoV-2 virions or synthetic gapd DNA) will be added, and one ‘no template’ negative control reaction. Increase the master mix amount proportionally if more replicates of each reaction are desired.

   Ensure that all individual reagents, especially those that are thawed from −20 °C, such as the reaction buffer and dNTPs, are uniformly mixed before use.

   For diagnostic applications of these two assays involving direct analysis of crude samples, such as heated human saliva, prepare a pair each of SARS-CoV-2 and gapd LAMP-OSD reactions and add the sample to one reaction tube (test) of each pair and an equal amount of water to the other (negative control). You will also prepare a third ‘no primer’ reaction tube (sample autofluorescence control) for each assay that will comprise all LAMP-OSD reaction components except LAMP primers. You will add the same amount of sample to these ‘no primer’ reactions in order to rule out any possibility of false positives due to sample autofluorescence.

   LAMP-OSD reaction compositions for detection of SARS-CoV-2 RNA and human gapd DNA provided in Tables 3 and 4 may be used as general starting points that may need further optimization when developing LAMP-OSD assays for other DNA or RNA targets.

2. Rehydrate the desired number of Br512 Mut235 cellular reagent tubes, each containing about 2 × 10⁸ dried bacteria (see Basic Protocol 1), with 30 μL of water by pipetting gently up and down.

   For performing five reactions (four containing different template amounts and one ‘no-template’ control) for each LAMP-OSD assay type, a total of 2 × 10⁸ dried bacteria will be required. Therefore, rehydration of two tubes of Br512 Mut235 DNA polymerase cellular reagents containing 2 × 10⁸ dried bacteria in each tube should be sufficient.

   The volume of water used to rehydrate the cellular reagents may be adjusted to match the starting volume of the bacterial suspension that was added to the tube during preparation of the cellular reagents (and noted on the cellular reagent container; see Basic Protocol 1).

   Cellular reagents should be rehydrated immediately before use. Do not store rehydrated cellular reagents for later use.

3. Incubate the rehydrated Br512 Mut235 cellular reagents at 65 °C for 30 min.

4. Briefly spin down the heated cellular reagent suspension to collect any condensates, and pipet gently up and down before transferring the desired volume into the pre-assembled LAMP-OSD reaction master mixes.

   For most LAMP-OSD assays, around 2 × 10⁷ dried bacteria of Br512 Mut235 cellular reagents prepared after overnight induction of protein expression from a tetracycline-regulated T7 promoter perform well. Therefore, use 1/10^th of the volume of the rehydrated 2 × 10⁸ dried bacteria (3 μL from a 30 μL rehydration volume) for each LAMP reaction in order to achieve 2 × 10⁷ dried bacteria/reaction.

   Once the cellular reagents are added to the LAMP-OSD reactions, the reactions should be processed for amplification and readout as quickly as possible. Do not store fully assembled assays for long durations.

5. Uniformly mix the LAMP-OSD master mix by gently vortexing or pipetting up and down and transfer the desired volume into individual thin-walled 0.2-mL tubes or 96-well plates.

   Typically, LAMP-OSD reaction is performed in a total volume of 25 μL that includes 2 to 5 μL of the template.

   Ensure that the LAMP-OSD reaction tubes or plates have no auto-fluorescence in the fluorescein range (or in the emission range of your fluorophore of choice) that could interfere with assay readout by visually inspecting empty tubes/plates in a transilluminator, gel doc, or DIY fluorescence visualization device (see Support Protocol 2).

6. Prepare the ‘no template’ negative controls for assessing amplification specificity by adding the required volume (usually 2–5 μL; the same volume should be used later in Step 8 when adding templates to positive control reaction tubes) of a non-specific template or water to the proper LAMP-OSD reactions and then cap or cover these reactions.

7. Prepare the LAMP templates as desired and, if necessary, dilute them in TE 10:0.1 buffer or water.

   For the SARS-CoV-2 assays, thaw and dilute the concentrated virion stock so as to add ≥500 virions/reaction to the four positive control assays. In a typical assay validation test, the four LAMP-OSD reactions may be seeded with different amounts of virions, such as 500,000, 50,000, 5000, and 500 virions/reaction, to prepare a curve. For the gapd LAMP-OSD assay, thaw and dilute the double stranded DNA templates so as to add ≥600 template copies/reaction to the four positive control assays. In a typical test, the four assays may be comprised of 600,000, 60,000, 6,000, and 600 template copies/reaction, to prepare a curve.

   For LAMP-OSD assays of other DNA or RNA targets, template copies used may vary depending on the detection limit of that assay. Typically, it may range from a few tens to several hundreds of thousands of copies per reaction.

   Use at least 2 μL as the template transfer volume in order to minimize variation due to pipetting errors.

8. Add the required amount of template to the proper LAMP-OSD reactions (excluding the ‘no template’ controls) and incubate all the reactions immediately at 65 °C for the desired duration.

   Unless the time-to-result of the LAMP-OSD assay for the template amounts being tested is known to be shorter, incubate the LAMP-OSD assays at 65 °C for 60 to 90 min.

9. Following completion of amplification, record a ‘yes/no’ readout of the assays by visually observing endpoint OSD fluorescence using a blue light transilluminator or gel doc used for observing DNA on ethidium bromide gels, or by using a DIY fluorescence visualization instrument (see Support Protocol 2).

   For some cellular reagent LAMP-OSD assays, such as the SARS-CoV-2 multiplex assay (see Table 3), the endpoint OSD signal may be enhanced by incubating the assays, post-amplification, at 95 °C for 1 min and then quick cooling them by exposure to ambient room temperature, ice, or 4 °C. Figure 7 shows an image of endpoint OSD fluorescence in SARS-CoV-2 multiplex LAMP-OSD assays performed using the Br512 Mut235 engineered DNA polymerase (Paik et al., 2021) cellular reagents and SARS-CoV-2 virions.

Figure 7. SARS-CoV-2 detection using multiplex LAMP-OSD assays and engineered Br512 Mut235 DNA polymerase cellular reagents.

Indicated copies of SARS-CoV-2 virions (determined from the droplet digital PCR genome equivalents/mL reported in the certificate of analysis by the vendor) were amplified in multiplex LAMP-OSD assays containing primers and OSD probes specific to regions in the viral N and ORF1ab sequences (see Table 3). Amplification was performed for 90 min using Br512 Mut235 DNA polymerase (Paik et al., 2021) cellular reagents followed by heating of the reactions at 95 °C for 1 min and subsequent readout of endpoint OSD fluorescence at room temperature. Images of endpoint OSD fluorescence acquired using a ChemiDoc gel imaging system (Bio-Rad, Hercules, CA, USA) are depicted.

SUPPORT PROTOCOL 2: A DIY FLUORESCENCE VISUALIZATION DEVICE

The following protocol describes how to build a DIY device for excitation of fluorescein (or other fluorophores whose absorption and emission wavelengths are similar to that of fluorescein) and visual observation of the emitted fluorescence. This device allows visual yes/no readout of LAMP-OSD assays (Basic Protocol 3) by enabling endpoint observation of accumulated OSD fluorescence directly by visual inspection or via an image captured using a cellphone camera. This protocol would facilitate application of LAMP-OSD in settings that lack other devices for detecting fluorescence, such as fluorimeters, blue light transilluminators, or gel doc systems. The device comprises a small dark housing box fitted with four blue 470 nm wavelength light emitting diodes (LED) on the bottom and a holder for four 0.2-mL tubes centered directly above the LEDs. A removable sliding lid allows LAMP-OSD reaction tubes to be loaded into the box. An observation window on the side is fitted with layers of inexpensive >500 nm bandpass gel filters that enable visual inspection of emitted fluorescence. A detailed list of parts and build documentation can be found at https://github.com/Jose-4625/LAMP-BOX-ECO/tree/master/VisBox.

While the protocol describes the use of 3D printer for building the body of the housing box, and employs soldering, breadboards, and power supply for building the electrical circuit, these can be readily substituted with alternative and/or lower-resource construction methods. For instance, for conducting initial feasibility tests and device design, we built our prototype devices by using small cardboard containers as the housing boxes and by attaching the LEDs, resistors, and connecting wires to them using electrical tape. The positive and negative wires of the units were connected to a 9V battery directly or via a simple battery holder to draw power.

Materials:

• Fluorescein (Sigma, cat. No. 46955, or equivalent)

• 3D-printed components for building the housing box (Figure 8):

  • U-piece (Upiece.stl)
  • Circuit tray (lampBOX circuit tray v1.stl)
  • Slider (slider.stl)
  • VisBox Body (VisBox.stl)
  • VisBox Top (VisBox.stl)
  • Components may be printed by accessing the stereolithography (STL) files listed within parenthesis next to each component from https://github.com/Jose-4625/LAMP-BOX-ECO/tree/master/VisBox. 3D-printed parts may be sourced from commercial vendors such as makexyz.com and printathing.com.

• 5V/12V breadboard power supply

• Small piece of insulated wire

• 4 × 100 ohm Resistors (Semtech Vitnette, item No. 100ohm100, or equivalent)

• 4 × Clear Blue LEDs (Microtivity, item No. IL041, or equivalent)

• 6 cm × 8 cm prototyping printed circuit board (PCB)

• Light filter gel pieces (Lee Filters, items 021 Gold Amber, 158 Deep Orange, 179 Chrome Orange, 767 Oklahoma Yellow or equivalent >500 nm bandpass filters)

• Sticky tape

• Soldering iron kit

Figure 8. Parts and assembly of a DIY fluorescence visualization device.

(A-D) Schematics for 3D printed parts designed to build the housing box – VisBox Body and VisBox Top (A), U-piece (B), Circuit tray (C), and Slider (D). I Circuit design. (F) Top view of the printed circuit board with attached power supply and LEDs. (G) Bottom view of the printed circuit board showing resistor connections. (H) Visualization box with LAMP-OSD tubes positioned for readout. (I) Picture (obtained using a cell phone) of endpoint OSD fluorescence in a LAMP-OSD test captured through the viewport of the fluorescence visualization box. Tube 1: ‘no template’ negative control; Tube 2: positive control tube with 10,000 template copies; Tube 3: positive control tube with 100,000 template copies.

Protocol steps:

1. Position the breadboard power supply at one of the short edges of the PCB such that the long edge of the breadboard lines up with the short edge of the PCB.

   Adjust the power supply position such that the pins at one of its short edges are aligned to the third row of holes in the nearest edge of the PCB.

2. Press the breadboard power supply into the PCB perforations.

   The PCB perforation spacing may be slightly different from the power supply pin spacing causing the bottom pins to bend slightly.

   Take note of the positions of the ground pins and the positive pins on the bottom of the power supply; pins of the same polarity should be aligned.

3. Solder the power supply pins that are farthest from the LEDs to the PCB (See bottom view in Figure 8).

   Ensure that the power supply is securely soldered to the PCB.

4. Place the longer (positive) leg of an LED in a PCB hole that is at least 5 holes away and in line with a positive power supply pin. Feed the short (negative) leg through the hole immediately underneath.

   Ensure that the LED is placed on the same side as the power supply. See Figure 8 for top view.

5. Skip three holes on the PCB and place another LED in line with the first.

6. Skip two holes and place the third LED in line with the others.

7. Skip three holes and place the final LED in line with the others.

8. Fold the long LED legs that are aligned with the positive pin of the power supply towards the positive pin (See bottom view in Figure 8).

   The folded LED legs should overlap with each other such that they can be used to connect the LEDs.

9. Solder the overlapping long legs of the LEDs and solder the LED leg closest to the power supply to the positive pin of the power supply.

10. Fold the short leg of the LEDs into small loops and feed them back to the power supply side of the PCB through the hole immediately underneath (See top view in Figure 8).

11. Connect a separate resistor to each LED by passing one resistor leg through the loop in the negative LED leads and soldering it in place. See bottom view in Figure 8. Cut off the extra material of the soldered resistor leg.

12. Bend the other legs of the resistors at 90° angles towards the power supply and solder them together at the overlaps.

13. Solder the resistor leg closest to the power supply to the negative pin of the power supply.

    If the last leg of the resistor is too short to reach the pin, add a piece of wire between the end of the resistor leg and the negative pin.

14. Assemble the 3D-printed parts by first placing the lower portion of the housing box on a flat surface and then placing the lid on top and attaching it to the U-piece with either glue or by soldering.

    Ensure that the slider piece can move freely in and out of the U-piece while the lid and U-piece are being attached.

15. Align the holes on the bottom of the housing box to the LEDs on the PCB and use tape to secure and level the housing box on the PCB.

16. Create the light filter for viewing fluorescence emission by layering pieces of yellow and orange gel filters and affixing this assembly over the viewport on the side of the housing box.

    Depending on the bandwidth and brightness of the LEDs used in the device, layers of 2 to 3 filter pieces can discriminate emitted fluorescence from incident LED light such that ‘no template’ negative control LAMP-OSD tubes (see Basic Protocol 3) appear dark when observed through the filter, while positive control LAMP-OSD reactions in which specific templates have been successfully amplified appear bright green (Figure 8I).

17. Set the power supply to 5V by toggling the jumper on the board and attach a power source to it using either a USB or a 12V barrel jack.

18. Verify performance of the fluorescence visualization box by observing fluorescence of 0, 50, 100, and 200 nM fluorescein in 25 μL volumes and adjust components, such as filter combinations, LEDs, and their distance from the bottom of the sample tubes, to optimize the signal-to-noise ratio, such that samples lacking fluorescein appear dark while samples containing fluorescein appear bright green (Figure 8I).

REAGENTS AND SOLUTIONS:

Lysogeny broth (LB)

• 10 g Tryptone (Sigma, cat. No. T7293, or equivalent)

• 10 g NaCl (Sigma, cat. No. S9888, or equivalent)

• 5 g Yeast Extract (Sigma, cat. No. Y1625, or equivalent)

• 950 mL water

• Dissolve all components in 950 mL water and then adjust pH to 7.0 using 5 N NaOH. Adjust final volume to 1 L by adding water. Aliquot into glass bottles with screw cap lids. Sterilize by autoclaving at 15 psi for 20 min on the liquid cycle with slow exhaust.

• Store at room temperature for up to 6 months.

Agar-containing LB medium

• Prepare liquid LB medium and add 15 g/L of agar. Sterilize by autoclaving at 15 psi for 20 min on the liquid cycle with slow exhaust. Swirl to uniformly distribute the melted agar and allow medium to cool to 50 °C to 60 °C before adding selection agents, such as antibiotics. Then pour 20 to 25 mL of medium per 100 mm Petri plate. When medium has hardened, invert the plates and store them sealed in a plastic bag at 4 °C.

• Store at 4 °C for up to 6 months.

10X ThermoPol buffer (may also be purchased from New England Biolabs, cat. No. B9004S)

• 200 mM Tris-HCl, pH 8.8

• 100 mM (NH₄)₂SO₄

• 100 mM KCl

• 20 mM MgSO₄

• 1% Triton^® X-100

• pH 8.8 at 25°C

• Store at −20 °C for up to 12 months or, if obtained from the commercial vendor, up to the manufacturer’s recommended expiration date.

10X G6B buffer

• 600 mM Tris-HCl, pH 8.0 (Invitrogen, cat. No. AM9855G, or equivalent)

• 20 mM (NH₄)₂SO₄ (Sigma, cat. No. A4418, or equivalent)

• 400 mM KCl (Invitrogen, cat. No. AM9640G, or equivalent)

• 40 mM MgCl₂ (Invitrogen, cat. No. AM9530G, or equivalent)

• pH 8.0 at 25°C

• Store at −20 °C for up to 12 months.

10X G6D buffer

• 600 mM Tris-HCl, pH 8.0

• 20 mM (NH₄)₂SO₄

• 400 mM KCl

• 80 mM MgCl₂

• pH 8.0 at 25°C

• Store at −20 °C for up to 12 months.

10X Isothermal buffer (may also be purchased from New England Biolabs, cat. No. B0537S)

• 200 mM Tris-HCl, pH 8.8

• 100 mM (NH₄)₂SO₄

• 500 mM KCl

• 20 mM MgSO₄

• 1% Tween^® 20

• pH 8.8 at 25°C

• Store at −20 °C for up to 12 months or, if obtained from the commercial vendor, up to the manufacturer’s recommended expiration date.

Annealed OSD probe (Figure 6)

• 100 μM fluorophore-labeled long OSD strand in TE 10:0.1 buffer comprising 10 mM Tris, pH 7.5 and 0.1 mM EDTA (e.g., see Table 3)

• 100 μM quencher-labeled short OSD strand in TE 10:0.1 buffer (e.g., see Table 3)

• 10X Isothermal buffer

• In a 0.2-mL PCR tube, mix 1 μM fluorophore-labeled OSD strand with 1 μM to 5 μM of the quencher-labeled OSD strand in 1X Isothermal buffer in a total volume of 100 μL. Using a thermocycler, incubate the tube at 95 °C for 1 min and then slowly cool down to 25 °C at the rate of 0.1 °C/sec.

• Store at −20 °C for up to 6 months.

1X SDS-PAGE sample buffer

• 63 mM, Tris-HCl, pH6.8

• 0.1% (vol/vol) 2-Mercaptoethanol (Sigma, cat. No. M6250, or equivalent)

• 0.0005% (wt/vol) Bromophenol blue (Sigma, cat. No. 114391, or equivalent)

• 10% (vol/vol) Glycerol

• 2% (wt/vol) SDS (electrophoresis-grade) (Sigma, cat. No. L3771, or equivalent)

• pH 6.8 at 25 °C

• Store at 25 °C for up to 12 months.

1X PBS buffer, pH 7.4

• 137 mM NaCl

• 2.7 mM KCl

• 10 mM Na₂HPO₄

• 1.8 mM KH₂PO₄

• Prepare a 10X stock of the buffer and, if necessary, adjust pH using hydrochloric acid or sodium hydroxide. Sterilize by filtration through 0.22 μm filter or by autoclaving at 15 psi for 20 min in the liquid cycle.

• Store at room temperature for up to 12 months.

SOC medium

• 2% (wt/vol) Tryptone

• 0.5% (wt/vol) Yeast extract

• 10 mM NaCl

• 2.5 mM KCl

• 10 mM MgCl₂

• 10 mM MgSO₄

• 20 mM glucose

Dissolve all components, except glucose, in water and then adjust pH to 7.0 using 5 N NaOH. Sterilize by autoclaving at 15 psi for 20 min on the liquid cycle with slow exhaust. In the meantime, prepare a 1M glucose solution by dissolving 18 g of glucose in deionized water, adjusting to a final volume of 100 mL, and filter-sterilizing using a 0.22 μm filter. After the autoclaved medium has cooled to ≤60°C, add sterile 1 M glucose solution to a final concentration of 20 mM. Store at room temperature for up to 6 months.

1M IPTG solution

• Dissolve 2.383 g of IPTG powder in water and adjust the final volume to 10 mL. Sterilize by filtration through a 0.22 μm filter and distribute into 1 mL aliquots. Store at −20 °C for up to 1 month.

0.1mg/mL Anhydrotetracycline solution

• Dissolve 0.1 mg of anhydrotetracycline powder per 1 mL of 100% ethanol and store the solution in a dark container at −20 °C for up to 6 months.

COMMENTARY:

Background Information:

Production of purified protein reagents usually requires overexpression in large-volume (multiple liters) cultures of recombinant E. coli or other engineered organisms to produce the requisite amounts of a protein of interest, followed by separation from undesired culture components using multiple steps and chemical reagents for cell lysis and protein isolation and enrichment (Wingfield, 2014, 2015). Many purification steps need to be performed in cold conditions, and most proteins, upon purification, must be stored at −20 °C to −80 °C to remain functional. While purification can be simplified using engineered tags, ranging from short peptides (such as six contiguous histidine residues) to entire proteins (such as the 42 kDa maltose binding protein), removing these modifications following protein purification adds complexity to reagent preparation (Arnau, Lauritzen, Petersen, & Pedersen, 2006; Goh, Sobota, Ghadessy, & Nirantar, 2017; Guan & Chen, 2014). While simple and inexpensive methods for sample and nucleic acid preparation have been developed to facilitate low-resource applications (Graham, Dugast-Darzacq, Dailey, Darzacq, & Tjian, 2021), the burdens of infrastructure, expertise, and time needed to purify proteins, mean that protein reagents for molecular biology can be readily purified only in appropriately equipped laboratories (Graham et al., 2021) and are, at present, primarily prepared at an industrial scale in a limited number of facilities.

The use of dried protein-expressing E. coli K-12 directly as reagents stands as an alternative route to carrying out reactions, and for simplifying the overall process of performing molecular and synthetic biological manipulations (such as DNA synthesis, amplification, and assembly) and molecular diagnostics. Most conventional and engineered enzymes for molecular and synthetic biology and diagnostics can be readily adapted to this cellular reagent platform. Examples include DNA polymerases, such as Taq (Chien, Edgar, & Trela, 1976), KlenTaq (Barnes, 1994), Bst-LF (Phang, Teo, Lo, & Wong, 1995) and its engineered variant Br512 (Paik et al., 2021), Vent (Jannasch, Wirsen, Molyneaux, & Langworthy, 1992), and Phusion DNA polymerases (Uemori, Ishino, Toh, Asada, & Kato, 1993; Wang et al., 2004); reverse transcriptases (RT), such as MLV RT (Roth, Tanese, & Goff, 1985) and the thermostable engineered enzyme RTX (Ellefson et al., 2016); and ligases, such as T4 and T7 DNA ligases (Becker, Lyn, Gefter, & Hurwitz, 1967; Cozzarelli, Melechen, Jovin, & Kornberg, 1967). In most cases, these cellular reagents are rehydrated in water and simply used instead of their pure commercial counterparts, without altering other reaction components, procedures, or analytical methods. Protein reagents do not need to be extracted or purified from the cellular reagents prior to use. At most, in some applications, such as LAMP or Gibson assembly, rehydrated cellular reagents may need to be pre-heated at 65 °C to 75 °C for 10 to 30 min prior to using them.

Both thermostable and mesophilic proteins, such as MLV RT, have been used in the cellular reagent format without pre-processing. In most analytical applications, such as qPCR, reverse transcription qPCR, endpoint PCR analyzed by agarose gel electrophoresis, and LAMP read using fluorogenic OSD probes (Jiang et al., 2015),, the performance of cellular reagents is comparable to that of reactions carried out with purified counterparts, as measured by amplification efficiency, detection limit, and time to result (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018). Similarly, cellular reagents can also be used as synthetic biology tools for building new DNA constructs via multi-enzymatic assembly methods, such as Golden Gate and Gibson assembly (although current efficiencies may be suboptimal for the construction of large libraries (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018).

The reason why cellular reagents perform as well as they do is unclear. At high temperatures characteristic of some procedures, such as the 95 °C denaturation step in PCR and the 65 °C amplification temperature of LAMP, we believe that the dried bacteria undergo thermal lysis likely due to weakening of cell wall integrity. E. coli membrane phospholipids have been found to melt and changes in peptidoglycan layer have been observed, and most proteins are known to denature at such temperatures (Leuenberger et al., 2017; Mackey, Miles, Parsons, & Seymour, 1991; Membrillo-Hernandez, Nunez-de la Mora, del Rio-Albrechtsen, Camacho-Carranza, & Gomez-Eichelmann, 1995). Indeed, in isothermal oligonucleotide extension assays of fresh Taq DNA polymerase-expressing E. coli, which were not dried into cellular reagents, we found that while the enzymes were largely inaccessible when the reactions were incubated at 37 °C or even at 42 °C, enzyme activity was robust at ≥65 °C, indicating thermal lysis of bacteria and release of their enzyme load (Bhadra, Pothukuchy, et al., 2018). Interestingly, in these same experiments, when we analyzed the activity of Taq DNA polymerase cellular reagents prepared from a portion of the same protein-expressing cultures, we observed strong oligonucleotide extension activity even in reactions incubated at 37 °C. These observations suggest that cellular reagents might have reduced cell wall integrity that upon rehydration allows some of their contents, including the heterologous enzyme load, to leak out and interact with externally added reaction components, thereby facilitating reactions (such as reverse transcription) at low temperatures that typically do not induce lysis of intact bacteria. Although this mechanism is speculative, it has been reported that when bacterial cells are desiccated or dried in the absence of external protectants (such as starch, trehalose, or skimmed milk), their phospholipid membranes tend to become leaky, especially upon subsequent rehydration (Berninger, Gonzalez Lopez, Bejarano, Preininger, & Sessitsch, 2018; Potts, 1994). This is because, as water is removed, the Van der Waals interactions between adjacent phospholipids increase, thus raising their phase transition temperature (Potts, 1994). This causes the lipids to transition into gel phase and separate from portions of the membrane that are still in the liquid-crystal phase. As a result, membrane permeability greatly increases and proteins associated with the membrane aggregate as they are excluded from parts of the membrane (Billi & Potts, 2002). Subsequent rehydration of the dried bacteria causes further damaging phase changes in the membrane, leading to leakage.

The quality of cellular reagents can be assessed by functional tests of enzyme activity (e.g., measuring qPCR Cq values (Bustin et al., 2009)), and given uniformity of bacterial growth and induction conditions, reliable batch-to-batch values have so far been observed (Figure 5). This allows cellular reagents to be simply used in place of pure protein reagents in existing protocols (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018). While cellular reagents can also be used to successfully perform molecular diagnostic reactions such as qPCR or LAMP, metrics commonly applied to the quality control of pure diagnostic reagents, such as the absence of nucleases and other cellular contaminants, may either be more difficult to apply to cellular reagents or may prove not to be relevant.

The cellular reagents described herein are attractive because of their (i) robust functional performance, (ii) ease-of-use via simply substituting for pure reagents in existing protocols, and (iii) retention of function at ambient temperatures during production and extended storage, without the need for a cold chain. The true value of the method we describe, however, arises from the facile production pipeline that does not rely on protein purification or complex instrumentation. The standardized methods we have described should enable preparation and use of common molecular biology reagents using only a bacterial incubator and a microcentrifuge, two relatively inexpensive and widely available instruments that can also be easily ‘homemade’ with locally sourced parts and readily available building instructions. For instance, build instructions for an Arduino-based incubator that we have used for preparing cellular reagents are documented at https://github.com/FOSH-following-demand/Incubator (Bhadra, Nguyen, et al., 2021) while several groups have described the construction of low cost minicentrifuges (Byagathvalli, Pomerantz, Sinha, Standeven, & Bhamla, 2019; WareJoncas, Stewart, & Giannini, 2018). Overall, the cellular reagent platform thus reduces the infrastructure, expertise, time, and ultimate cost burdens associated with reagent production, storage, and distribution.

We envision that uptake of these and other lower-resource methods may provide entirely new opportunities for local and regional economies to directly benefit from the public health, research, educational, and other bounties that biotechnology enables. Starter expression constructs can be readily engineered and then distributed via plasmid repositories or sequence databases. The protocols presented should thus enable production of commonly used molecular biology proteins as cellular reagents with minimum cost, infrastructure, and expertise, even in low-resource settings, and it should now prove possible to sustainably make or adapt cellular reagents according to local priorities, without the need to depend on globalized supply chains or goods that can only be produced by multinational companies or governments. Eventually, this process should feedback on itself given that cellular reagents themselves can be used for DNA synthesis and assembly.

The feasibility of rapidly producing large amounts of cellular reagents on site, even in remote locations, may cut expenses and response times to the point where they prove attractive for widespread public health programs for surveillance of epidemic or pandemic disease prevalence. Ultimately, diagnostic performance criteria, such as sensitivity, specificity, accuracy, repeatability, and limits of detection, are the most important factors for diagnostic success. Furthermore, given the desirability and proven diagnostic accuracy of tests containing impurities, such as those introduced when analyzing crude samples (Vogels et al., 2020), it is conceivable that cellular reagents might find regulatory acceptance for monitoring non-human hosts, such as mosquitoes and ticks for arthropod-borne diseases, or indirect human samples, such as sewage or contaminated water bodies for pathogens or human fecal indicators. Building on the success of such implementations in environmental testing, it eventually might become feasible to imagine applications of cellular reagents in medical testing, particularly during unprecedented emergencies or for large scale screenings.

Critical Parameters:

Overall, the key to preparing and using cellular reagents is standardization. Bacterial growth and protein induction conditions should be optimized beforehand or reproduced from previous publications, while processing of overexpressing strains into dried cellular reagents must be carried out according to the protocol presented here in order to ensure reliable and reproducible results.

It is good practice to use freshly-transformed bacteria for preparing cultures for protein induction, since the metabolic burden of plasmid maintenance and protein expression, along with plasmid segregational instability, might favor selection of bacteria containing mutated plasmids and reduced ability for high-level protein synthesis (Bentley, Mirjalili, Andersen, Davis, & Kompala, 1990). Additionally, classic protein expression strains such as BL21(DE3) have high spontaneous DNA mutation rates (Csorgo, Feher, Timar, Blattner, & Posfai, 2012). The choice of bacterial growth medium can also influence protein yields. In most cases, well-known rich media, such as LB broth or 2X YT broth, or proprietary media such as Superior Broth^™, specifically formulated for high yield protein production, yield good quality cellular reagents.

To guide induction and subsequent preparation of cellular reagents, culture density should be measured accurately. While logarithmic growth phase cultures at A₆₀₀ of 0.5 to 0.7 can be measured directly, denser cultures, such as those that have been induced for protein expression for 3 h or longer may need to be diluted before measuring their absorbance. If using a spectrophotometer for measuring absorbance, it should be calibrated using the appropriate blank, such as the culture medium or the diluent, in each case. If using McFarland turbidity standards, ensure that the standards are shaken well and uniformly turbid. If the standards are stored for prolonged use, ensure that they are tightly capped and check for uniform turbidity and lack of evaporation prior to each use.

Once cells have reached the desired density, perhaps the most critical parameter for successfully preparing cellular reagents is control over induction. For most nontoxic proteins, strong E. coli- or oT7 phage promoter-driven transcription of the gene of interest from a medium-to-high copy plasmid works well. Toxic protein expression may need to be performed using low-copy plasmids and engineered E. coli strains (e.g., T7 Express lysY, NEB, cat. No. C3010I) designed to reduce leaky expression. Most commonly used protein expression systems can, in turn, be regulated by transcription repressors, such as LacI or TetR, that require specific chemical additive(s) to induce protein expression. The concentration of the chemical inducer added to the bacterial culture should be optimized for the desired strength of induction for the bacterial strain and the plasmid expression system being used to prepare cellular reagents. Briefly, several replicate cultures of the transformed bacteria should be prepared and while allowing some cultures to continue growing without the addition of the inducer, the other cultures should be treated at the same growth stage (as determined by culture density measurements) with different amounts of the inducer. After allowing all cultures to grow for the same post-induction duration, their level of protein expressed should be compared using methods such as SDS-PAGE analysis (Support Protocol 1). Attention should also be paid to the timing of induction, both with respect to when the inducer is added to the culture and the duration of induction. For most protein expression systems, the inducer should be added to cultures at an A₆₀₀ ranging from 0.5 to 0.7 to give optimal results. Inducing cultures too early or too late in their growth phase may lead to reduced protein yield and, consequently, less active cellular reagents. Induction duration often depends on the temperature of induction, as well. Inductions at 37 °C are usually performed for a short duration, typically for 3 to 4 h; prolonged induction of cultures at 37 °C can be deleterious for cellular reagent applications because overgrowth and nutrient depletion might result in cell death, protein aggregation, and protein degradation (Donovan, Robinson, & Glick, 1996). Proteins that need to be expressed at lower temperatures to improve stability or minimize toxicity usually need ~18 to 20 h of induction. Finally, care should be taken to follow manufacturers’ guidelines for storage, preparation, and expiration date of inducers to ensure optimal protein induction.

Once cells are ready for processing, the recommended amount of bacterial suspension in a single aliquot prepared for drying is 20 to 35 μL per 0.2-mL tube. Smaller volumes may lead to more discrepancies in bacterial numbers per aliquot, while larger volumes may take longer to fully dry. Although dry cellular reagents are stable when stored at 37 °C for extended durations, moist bacteria kept at 37 °C for longer than 24 h may result in reduced activity of the resulting cellular reagents. While a desiccator is not required for storing dry cellular reagents, they should be tightly capped and stored in a closed container, such as a Tupperware box, in the presence of desiccants, ideally at 4 °C to 25 °C.

For the use of cellular reagents, rehydration should only be done immediately prior to addition in molecular reactions, and hydrated cells should not be stored for future use. Pipetting up and down is usually sufficient for rehydration; vigorous vortex mixing is not necessary. For most molecular biology applications, cellular reagents are either rehydrated using water and aliquoted into desired reactions, or they can be directly rehydrated in an appropriate amount of the reaction master mix. For some applications, such as LAMP-OSD or Gibson assembly, rehydrated cellular reagents need a brief thermal pretreatment at 65 °C or 75 °C before use. We surmise that this heat treatment likely helps the cellular reagent reaction by inactivating some of the endogenous DNases that might otherwise compromise the outcomes. In most protocols, cellular reagents are used at 2 × 10⁶ to 2 × 10⁷ dried bacteria/reaction. As described in Basic Protocol 1, this is an extrapolated number derived from the final A₆₀₀ value of the washed bacterial suspension used to prepare the cellular reagent and the approximation 0.5 A₆₀₀ = 5 × 10⁸ bacteria/mL. Consequently, if the cellular reagent preparation contains 2 ×10⁸ dried bacteria/tube, add water (usually 30–40 μL) and use 1/10^th of the rehydrated volume (3–4 μL) per reaction in order to achieve 2 × 10⁷ dried bacteria/reaction. The amount of cellular reagents that is required in a reaction usually depends on the strength and duration of protein induction performed during preparation of the cellular reagent. The amount used is also contingent on the type of reaction. For instance, a 20-μL Gibson assembly reaction usually contains 2 × 10⁷ Taq DNA ligase cellular reagents, but only 1 × 10⁵ and 1 × 10⁶ of similarly prepared T5 exonuclease and Taq DNA polymerase cellular reagents, respectively. Since both too many and too few cellular reagents in a reaction can be detrimental, users should experimentally determine the optimal amount of their cellular reagent preparations per application.

Cellular reagents are usually prepared by drying bacterial suspensions in 1X PBS. As such, and as an additional consideration, the cellular reagent preparation protocol may require substitution of PBS with a different buffer or water for reactions that are sensitive to one or more components of PBS, such as hydrolysis of phosphate monoesters catalyzed by alkaline phosphatase. This may, in turn, lead to alterations in drying times, the physical and functional properties of the resulting cellular reagents, and even rehydration conditions. Application conditions for such cellular reagents may also need to be re-optimized.

While cellular reagents can be used for a variety of molecular biology applications, their use in nucleic acid amplification assays is especially noteworthy, since, in most cases, their performance has been shown to be on par with that of pure proteins (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018). When performing nucleic acid amplification reactions such as PCR or LAMP, it is imperative to minimize sources of cross-contamination with templates. Therefore, in all procedures, before handling any templates, all other amplification reaction components should be assembled into master mixes and distributed into reaction tubes, preferably using barrier pipette tips. Also, the negative control reaction tubes should be prepared and tightly capped before handling any templates. While preparing templates or samples, test reaction tubes or plates should be kept covered with caps or a piece of clean paper, plastic, or cardboard to minimize aerosol contamination. If possible, a separate area and a separate set of pipettes with barrier tips should be used for preparing templates and their dilutions for amplification. If preparing template dilutions, small volumes with respect to the container (for example, 50 to 200 μL volume in a 1.5-mL tube) should be used, such that during mixing of this volume of liquid by vortexing or flicking, the templates are less likely to reach the tube lids, thereby reducing the chance for aerosolized contamination when the tube is subsequently opened. In addition, all reagents and templates should be spun down prior to opening the tubes, in order to further minimize contamination. After use, pipettes, work area, and ancillary equipment, such as tube racks, should be cleaned with 70% ethanol (or equivalent) and gloves discarded or washed after handling templates. Lastly, it should be noted that since cellular reagents retain bacterial genomic DNA, they are not suitable for diagnostic applications intended to detect sequences that may also be found in the laboratory E. coli strains used to prepare cellular reagents.

While there will be a tendency to be suspicious of a ‘crude’ cellular reagent, the primary source of problems in amplification reactions remains amplicon cross-contamination rather than the reagents. Contamination of LAMP reactions with preformed LAMP amplicons can be especially difficult to overcome, because they can prime their own continuous amplification and rapidly consume all reaction resources, thereby suppressing true signals. Therefore, to ensure confidence in the reactions, it is generally advisable to avoid opening the LAMP reaction tubes during or after the amplification is complete. If tubes containing LAMP amplicons must be opened, such as for examination by agarose gel electrophoresis, it should ideally be done in a space set well apart from where LAMP reactions are assembled. In addition, a dedicated set of pipettes that is not used to assemble LAMP reactions is preferable for handling LAMP amplicons in order to further minimize chances of amplicon contamination. Several readout methods that do not require opening of LAMP reaction tubes have been reported. Some use non-specific readout tools, such as color changing metal-sensitive indicators or pH-sensitive dyes (Tanner et al., 2015; Tomita et al., 2008); others use sequence-specific probes, such as OSD probes (Jiang et al., 2015), or fluorophore-labeled primers (Tanner, Zhang, & Evans, 2012). Given the propensity of continuous amplification methods such as LAMP to produce spurious amplicons, a probe-based readout of LAMP reactions carried out with cellular reagents is likely to give the best results.

Troubleshooting:

Table 5 lists common problems with the protocols, their causes, and potential solutions.

Table 5.

Troubleshooting Guide for preparation and use of cellular reagents.

Problem	Possible Cause	Solution
No / poor protein expression	Degraded inducer	Prepare fresh inducer solution or if appropriate, add measured amount of powder directly to the culture.
	Culture over- or undergrown at the time of induction	Prepare a fresh culture for induction.
	Mutated plasmid or protein expression cassette	Sequence the protein expression plasmid and reconstruct or reacquire correct plasmid. Use fresh transformants for preparing cellular reagents.
	Suboptimal induction conditions	Compare the level of expression of your protein in induced bacteria by comparing with duplicate uninduced cultures using SDS-PAGE analysis. Systematically test the effect of different induction conditions on protein levels. For example, initiate induction at culture densities ranging from A600 values of 0.5 to 1.0, use different amounts of inducer, and vary temperature and duration of induction (3–4 h at 37 °C to overnight at 18–30°C). Repeat SDS-PAGE analysis to assess changes in levels of protein expression. In depth guidance on optimizing heterologous protein expression in E. coli may be found in (Francis & Page, 2010).
Wet cellular reagents after 24 h at 37 °C	Exposure of desiccant to moisture	Use moisture-indicating desiccants, such as indicting Drierite™, to visually assess that the desiccant is dry. Use a new batch of desiccants or regenerate dry desiccants, such as calcium sulfate granules, by spreading them in a single layer and incubating in an oven set at 210° C for 1 hour. Tightly cap the desiccant container in which the cellular reagents are being dried.
	Too much bacterial suspension added to each tube	Do not dry more than 35 μL of bacterial suspension per 0.2-mL tube.
	Insufficient desiccant amount	Do not overload desiccant containers with cellular reagents. For example, place three to four 0.2-mL × 8-tube strips in about 250 g of calcium sulfate desiccant.
	Insufficient opening in the tubes for efficient evaporation	Leave the tube caps entirely open. For capped tubes, make a hole through the cap using an 18-gauge syringe needle.
	Desiccant container left ajar	After placing the cellular reagents inside the desiccant container, tightly close the container lid.
Inactive or poorly active cellular reagents	Poor protein induction during preparation	Remake cellular reagents and check level of induced protein by performing SDS-PAGE gel electrophoresis.
	Too many or too few cellular reagents used in the reaction	Optimize the amount of cellular reagents used in the reaction.
	Nuclease activity in cellular reagents	For thermotolerant and thermostable cellular reagents, prepare them in endonuclease-deficient strains and/or heat-treat hydrated cellular reagents prior to use at 65 °C to 95 °C for 5 to 30 min. For thermolabile cellular reagents, prepare them in endonuclease-deficient strains, such as DH5α.

Understanding Results:

E. coli transformed with a protein expression plasmid and plated on an agar growth medium containing appropriate antibiotics should yield colonies within 18 h to 24 h of incubation at 30 °C to 37 °C. Single colonies cultured overnight in rich broth medium containing selective antibiotics are usually dense enough to appear almost opaque. Most 1:200 aerobic E. coli sub-cultures transformed with a medium- to high-copy plasmid should usually reach ~0.5 A₆₀₀ within 2 h to 3 h of growth at 37 °C, especially BL21 strains. Following completion of induction, cultures typically reach an A₆₀₀ value >1.0. Suspensions containing washed, induced bacteria in 1X PBS at a concentration of 6.0 to 10.0 A₆₀₀ appear translucent to opaque, even when dispensed in small volumes inside 0.2-mL tubes (Figure 2). Upon drying at 37 °C in the presence of desiccants, the cellular reagents appear as a yellowish and powdery coating on the tubes. Freeze-drying in a lyophilizer produces a white powdery appearance (Figures 2 and 3). When cellular reagents that have been dried at 37 °C are rehydrated, the suspension appears turbid (albeit less turbid than the bacterial suspension prior to drying). When lyophilized cellular reagents are rehydrated, they display even less turbidity compared to the original or dried bacterial suspensions.

If SDS-PAGE is used for assessing protein expression, the protein of interest can usually be visualized on a Coomassie-stained gel loaded with the equivalent of 2–10 × 10⁸ induced bacteria (Figure 9). Similar numbers of uninduced bacteria should either not display the protein of interest or, in case of leaky expression, may display a faint band. Bacteria that have not been transformed with the plasmid of interest should be included in the SDS-PAGE analysis as a negative control, for comparison.

Figure 9. Example SDS-PAGE analysis of bacteria expressing a protein of interest.

Lysates were obtained from ~2 × 10⁸ bacteria that were either induced for 3 h (lane 1) or kept uninduced (lane 2) and were analyzed by SDS-PAGE. An image of the Coomassie-stained gel is depicted. The protein band of interest is indicated by an arrow.

In functional tests, the performance of optimized amounts of many cellular reagents of proteins, such as DNA polymerases, has been shown to be comparable to that of their purified counterparts. For instance, cellular reagents for Taq or Bst DNA polymerases demonstrate similar limits of detection and time-to-result in qPCR and LAMP-OSD assays (Bhadra, Pothukuchy, et al., 2018). When stored properly, this level of performance can usually be maintained for several months. Batch-to-batch performance of cellular reagents is also consistent and reproducible when similar conditions are used for preparation; for instance, the same bacterial strain and expression system, the same growth conditions, and the same timing of induction (Figure 5). Other cellular reagent-driven reactions, however, may demonstrate lower efficiency than with their pure counterparts. For instance, cellular reagent-mediated DNA assembly is less efficient than commercial pure enzyme master mixes (Bhadra, Pothukuchy, et al., 2018). That said, sufficient numbers of correct clones, as verified by sequencing, can likely be obtained to support using cellular reagents for the assembly of DNA constructs.

Time Considerations:

When starting with an optimized protein expression system, the cellular reagent preparation described in Basic Protocol 1 can be completed in 4–5 consecutive days. Preparation of the McFarland standard as described in Alternate Protocol 2 should take about one hour, while SDS-PAGE analysis followed by Coomassie protein staining as described in Support Protocol 1 should take at most 2–3 h to run the samples on a precast gel, and from 1–2 h hours to over a day to stain and visualize the proteins. Reactions performed with cellular reagents should take similar amounts of time to set up and obtain results as the same reactions using pure protein reagents. For instance, PCR or LAMP amplification assays performed using ready-to-use templates and cellular reagents, as described in Basic Protocol 2 and Basic Protocol 3, respectively, can be completed within 3 to 4 h depending on user familiarity with the procedure and the duration of amplification. The DIY instrument described in Support Protocol 2 can be built within a day or two if all parts and instruments are available.

DATA AVAILABILITY STATEMENT:

All new data that support this protocol are included in the article. All previous data supporting this protocol are openly available (Bhadra, Nguyen, et al., 2021; Bhadra, Pothukuchy, et al., 2018)].