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Use of Microfluidics for Directed Evolution of Biocatalysts and Materials in Droplets and Microgels

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Submitted:

23 February 2024

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26 February 2024

Withdrawn:

15 March 2024

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Abstract
Microfluidics is used for droplet-based biochemical and biological assays in protein and metabolic engineering for designing novel biocatalysts. These systems are based on lab-on-chip and glass capillary devices that enable the encapsulation of organic and aqueous phase materials. Recently, together with lab-on-chip microfluidics, novel devices based on modular LEGO microfluidics have been developed that can be used to produce complex droplets and direct the evolution of proteins, cells, and materials in liquid droplets or microbeads. In vitro, compartmentalization of single genes or cells can be done in liquid microdroplets, microbeads, or microcapsules using microfluidic methods. Most often, substrates and detection systems based on absorbance, electrochemical measurements, fluorescence, mass spectrometry, and Raman spectroscopy can screen for more active protein variants. Efforts have also recently been made towards integrating microfluidics and flow cytometry, which are incompatible due to the use of different emulsion systems. Still, there are trials to achieve full automation and integration of these complex platforms. A novel approach emerges, where beads or capsules can be a substrate for compartmentalized enzymes, and cells, and this review will also tackle this most recent approach and the possibility of using it for the directed evolution and engineering of biocatalysts and (bio)materials.
Keywords: 
Subject: Biology and Life Sciences  -   Biochemistry and Molecular Biology

1. Introduction

The lack of rapid high-throughput screening (HTS) systems has been a major bottleneck in creating biocatalysts via directed evolution [1]. With advanced robotics, microtiter plates (MTPs) can screen libraries of up to 105 clones per day [2] which is ~1/s, albeit at a high cost. Screening of single cells by fluorescence-activated cell sorting (FACS) allows analysis of up to 50,000 cells per second. Still, it is only possible under two conditions: (a) the produced enzyme stays in the cytoplasm and the substrate is able to diffuse into the cell while the fluorescent reaction product is trapped inside the cell or (b) the enzyme is displayed at the cell surface and the fluorescent reaction product is also attached on the cell surface. Both scenarios enable the colocalization of genotype (catalytic activity indicated by fluorescence) and phenotype (the enzyme encoding gene).
Emulsion droplets or particles can serve as miniaturized reaction vessels that can link phenotype to genotype even if the fluorescent reaction product is released from the cells or separated from the enzyme encoding gene. In this approach droplet boundary insures the genotype/phenotype linkage. However, the interface between the aqueous and oil phases is not completely impermeable as the plastic or glass surface of test tubes and multi-well plates and small molecules can diffuse between the droplets. It can be minimized using suitable oils and surfactants or jellifying droplets to reduce the diffusion rate within the aqueous compartments. In this review, we discuss recent advances in using microfluidics for HTS screening of biocatalysts and biomaterials. We review the advantages of microfluidics compared to traditional bulk emulsification methods and various microfluidic strategies for performing both processes.

1.1. Advantages of droplet-based screening methods compared to multi-well plates

Due to the micrometer dimension of droplet compartments, reaction volumes can be reduced from the microliter to the picoliter range, which allows screening at the level of single cells or even genes. For example, a droplet with a diameter of 10 µm has a volume of 0.5 pL which represents a volume reduction >3×108 compared to the regular 96-well plates with a ∼200 μL volume. This is necessary because the possible combinations of amino acids – even in a focused protein library – easily exceed the screening capacity (e.g., a library in which only 5 residues are fully randomized almost matches the throughput of droplet microfluidics [3]. A comparison between microtiter plates, FACS and droplet-based screening methods was reviewed recently [4].

1.2. Advantages of microfluidics in HTS systems

Microfluidics allows high screening rates in small volumes without capillarity or evaporation problems, high encapsulation efficiency of compartmentalized species due to small shear forces, integration of multiple unit operations on a single chip, flexible and versatile channel geometries that can be designed quickly using computer-aided design (CAD) software and replicated with high precision during microfabrication, high control over compartment size and its internal structure and composition, continuous production, automated liquid handling, possibility of connecting several chips in a modular system, minimal consumption of reagents and plasticware (tubes, plates, and pipette tips), and visual control over the process since microfluidic channels are optically accessible. In addition, production of monodisperse emulsions in microfluidic channels allows a more stringent quantification of the reaction product based on the optical readout [3]. Furthermore, droplets can be individually manipulated (created, fused, split, incubated, probed, and sorted) with high precision and the timing of mixing, lysis, reaction, incubation, and other steps can be precisely governed by the geometry of microfluidic channels and fluid flowrates.

1.3. Bulk emulsification versus droplet microfluidics

Bulk emulsification methods are rapid, can generate 1010 droplets in less than 5 min, and require simple and cheap equipment. However, bulk emulsification is typically a batch process and the droplets formed using these methods show a wide variation in size. With bulk emulsion, a single gene encapsulation is difficult to achieve since each twofold change in droplet radius corresponds to an eightfold change in volume, so large droplets dominate the total volume and are much more likely to be occupied by multiple cells / genes.
In microfluidic devices droplets are produced in a continuous flow manner at rates lower than those of bulk emulsion but still >106/h [5]. More importantly, droplets are monodisperse with a typical variation in size of <3% which satisfies the NIST definition of monodispersity (“a particle distribution may be considered monodisperse if at least 90% of the distribution lies withing 5% of the median size”). Moreover, double emulsion droplets with core-shell morphology can easily be produced in microfluidic chips, which makes it possible to screen droplets using FACS. Using bulk emulsification methods, typically each outer droplet consists of many inner droplets and such inner droplets cannot be screened one by one because their projected areas overlap as they travel along the channel during screening. On the other hand, microfluidic devices require highly specialized equipment to be fabricated and skillful operators. Finally, microfluidic devices are often disposable due to wetting and clogging problems, so usually a new chip must be manufactured after each experiment.

2. Fabrication methods for microfluidic chips

2.1. Soft lithography

Microfluidic chips can be made by soft lithography in polydimethylsiloxane (PDMS) using the protocol developed by Xia and Whitesides [6]. The soft lithography process involves two main steps: creating the master mold by photolithography and forming the polymeric chip by molding (Figure 1 a). To create the master mold, a layer of negative photoresist (SU-8) solution is applied onto a silicon wafer by spin-coating, followed by soft baking to evaporate the solvent and create a dry film. Ultraviolet light is then passed through a photomask to pattern the photoresist that is subsequently “post baked” to complete the polymerization process started by UV light. The photoresist can also be patterned by applying the projected light or focused beam of charged particles (electrons or ions) directly onto the resin surface without the need for a mask, which is known as maskless (direct-write) lithography. After lithography, the unpolymerized photoresist is dissolved using a developer (propylene glycol monomethyl ether acetate). In the second step, liquid PDMS is poured into the master mold, baked to crosslink PDMS and form the polymerized device, and finally bonded onto glass (or another PDMS surface) via oxygen plasma treatment to seal microfluidic channels and create a leak-proof device. The channels can be connected to pumps and pressure reservoirs via tubing attached to tube connectors that are inserted into holes made with biopsy punches. Compared to silicon and glass chips, PDMS chips can be fabricated rapidly once the master mold is available, but they suffer from deformation at high pressures and swelling in contact with organic solvents [7].

2.2. Photolithography and etching

Glass chips can be manufactured by photolithography (Figure 2b) and isotropic wet etching using hydrofluoric acid (HF) (Figure 1d) [8]. Dry etching and laser ablation can be used to create channels with sharp edges without mask undercut, albeit at smaller etch rates [9]. Glass chips can be bought “off-the-shelf” from Dolomite Microfluidics. Glass is more rigid and resistant to organic solvents than PDMS, has superior optical properties and high thermal resistance, making glass chips reusable but more expensive to fabricate. Shallow structures in single crystal silicon and quartz chips are patterned using anisotropic wet etching, e.g. by potassium hydroxide (KOH) (Figure 1c) or tetramethylammonium hydroxide (TMAH) [10], while reactive dry etching (RIE) and deep reactive ion etching (DRIE) (Figure 1 e) are used to etch deep narrow structures. The main problems of silicon chips are a time-consuming and expensive fabrication process, and fragility resulting from their inherent brittleness.

2.3. Hot embossing and injection molding

Chips from thermoplastics such as poly(methyl methacrylate) (PMMA) and cyclic olefin copolymer (COP) can be fabricated using hot embossing (Figure 1 f) and injection molding. Hot embossing is based on pressing silicon mold into a softened polymer heated above the glass transition temperature. Once the polymer has conformed to the shape of the mold, it is cooled below the glass transition temperature so that it is sufficiently hard to be separated from the mold. Injection molding is the process by which a softened polymer is introduced into a mold cavity under high pressure. Injection molding is more suitable for large scale production of chips, but it becomes costly for small-scale production, due to the need for expensive micromachining tools.

2.4. 3D printing

3D printing has become increasingly popular in recent years [11] due to the decreasing costs of 3D printers, an increase in printing resolution, and the ability to create true 3D channels. Fused deposition modeling (FDM) 3D printing involves injection of a hot polymer melt through a nozzle onto an XYZ stage to build up a three-dimensional structure layer-by-layer (Figure 1 g). The maximum resolution that can be currently achieved using this technique is about 50 μm. SLA (stereolithography) and DLP (digital light processing) 3D printers build up material through the polymerization of a liquid photopolymer using a guided laser beam (SLA) or a projector (DLP). The crucial difference is that DLP 3D printers use a digital projector screen to flash an image of a layer across the entire platform, curing all points at the same horizontal position simultaneously [12].

2.5. Laser ablation

Laser ablation is a process in which a laser beam removes material through vaporization. It can be used for the maskless generation of microfluidic patterns on substrates such as epoxy resin [13], polymethylmethacrylate (PMMA), PVC, glass [14,15], etc. The laser system can also be used for the drilling of inlet/outlet ports and the bonding of two plates together to enclose the laser-generated patterns from the top (Figure 1 h).

2.6. Micro-cutting

Micro-cutting includes drilling and milling and the micro-tools are fabricated by other methods of smaller resolution. Drill diameters less than 25 µm are available and the tool speed and position are usually controlled by CNC. Micro-milling can easily manufacture 3D structures of high aspect ratios with inclined angles on side walls [16]. There also exists a wide array of machinable materials for micro-milling, such as metals, polymers, composites, and ceramics. Stainless steel is particularly advantageous and cannot be machined with traditional MEMS approaches (Figure 1 i).

2.7. Glass capillary pulling

Glass capillary microfluidics is based on using coaxial assemblies of borosilicate glass capillaries. Typically, inner capillaries with a tapered end are prepared by pulling commercially available capillaries with a millimeter scale diameter to reduce their inlet diameter to several tens or several hundreds of micrometers [17].

3. Modular microfluidics

Traditional microfluidics is based on using monolithic chips that can perform multiple operations. However, monolithic chips lack operational and structural flexibility and are difficult to fabricate, operate, and reconfigure. The modular microfluidics concept is based on using standardized building blocks for individual fluidic operations that can be designed, fabricated, and tested separately and easily assembled into a customized, re-configured microfluidic platform [18,19,20]. To form a platform, individual pre-fabricated chips need to be connected in such a manner as to prevent leakage and ensure the smooth operation of the whole platform. Microfluidic chips can be connected using the LEGO stud-and-tube connection [18], the tubing connection [21], the capillary connection [22], the Luer connection [23], and the plasma connection [24].

3.1. LEGO connection

In this approach, the building blocks are stackable 2  ×  2 Lego-like® bricks made of PDMS using soft lithography, that can be reversibly assembled on a Lego® base plate [18]. Microfluidic channels with a cross-sectional width and height of 500 µm were imprinted on the building blocks' bottom and lateral surfaces. To form a 2D channel network, channels on the bottom surfaces of adjacent building blocks were connected at the block edges and the channels were sealed by the Lego® base plate or a building block's top surface. 3D channel networks were formed from channels that perpendicularly extended from bottom surface channels and were imprinted on the lateral surface of a building block. Lateral surface channels terminated at the top surface of the same building block and these channels were sealed by the lateral surface of an adjacent building block (Figure 2).
LEGO-like plastic blocks manufactured by CNC milling have been also used to connect two parts of the same microfluidic module [25]. The advantages of this approach are that the module can be easily dismanted for cleaning and reasssembled.

3.2. Tubing connection

In this approach, individual chips are connected using flexible plastic tubing inserted into inlet and outlet ports of two adjecent modules. This connection was used to connect an automated concentration gradient generator and a single-cell trapper array for capturing, culturing, and observing individual cells [26]. The same modular approach was used to construct a concentration gradient generator by connecting multiple chips with simple channel geometries via plastic tubes [27]. A concentration generator based on modular architecture can be used to produce complex and tunable gradient profiles by adding, subtracting, or replacing individual, standardized parts with simple channel patterns. In conventional approach based on using a single integrated gradient generator, each new design of gradient require another microfabrication.

3.3. Capillary connection

In this approach, individual CNC-milled chips are connected via glass capillaries screws, and O-rings [28]. In Figure 3, two CNC-milled modules are connected via inner capillary with two tapered ends and used to produce double emulsions with controllable number on inner droplets ranging from several to several hundreds [29].
To make a W1/O/W2 emulsion, inner droplets are generated by flow focusing of an inner water phase by an oil stream at the upstream tip of a long inner capillary connecting the two module, while the outer droplets are generated at the downstream tip of the same capillary by co-flow of the generated W1/O emulsion and the outer water phase, Figure x. Depending on the geometry of the capillaries and fluid flow rates, the number of encapsulated inner droplets can vary from two to over one hundred.

3.4. Luer connection

In this case, individual 3D-printed parts are connected via miniaturised Luer-Lock fittings [30]. Yuen developed a “Plug-and-Play” modular microfluidic system for biological and chemical applications composed of of a motherboard with interconnected grooves, H- and T-shaped microchannel inserts with two female Luer-Lock fittings connected with an internal straight channel, and chips with different functionalitities containing male luer lock fittings (Figure 4). The integrated system is assembled by placing microchannel inserts in a positioning groove of the motherboard and then plugging the chips into the microchannel inserts. Tubing was plugged into the microchannel inserts via a pipette tip which was cut to fit into the female Luer fitting.

3.5. Plasma connection

In this configuration, multiple PDMS chips were stacked together and bonded by plasma bonding. This approach was used to generate large-scale perfusable microvascular networks [31]. The culture medium can flow between the two chips via communication pores.

4. Integration of compartment manipulation, detection, and sorting methods in microfluidics and flow cytometry

In vitro compartmentalization (IVC) technologies are discovered by Tawfik and Griffiths. They involve the compartmentalization of genes into aqueous microdroplets dispersed in an inert oil [32]. The maintenance of the genotype-phenotype linkage in this system is achieved by the co-compartmentalization of single genes, the substrate for the enzymatic reaction being studied, and an in vitro transcription/translation solution for protein expression. Nowadays, this technology can be applied to compartmentalize cells or single genes in W/O emulsion droplets, W/O/W double emulsion droplets, microgels, giant liposomes, etc. Water-in-oil (W/O) emulsion droplets for directed evolution were first produced in a polydisperse system using traditional bulk emulsification devices: for single-cell experiments (in vivo compartmentalization) [33], or for single-enzyme experiments (in vitro compartmentalization) [34].
While the droplet boundary can largely restrict crosstalk, droplet sizes generated by bulk emulsification vary considerably and consequently the assay quality may differ from droplet to droplet, as differently sized droplets will contain different amounts of reagents. Nevertheless, polydisperse emulsions are still used today for protein engineering, especially if there is no need for quantitative measurement of the reaction product, i.e., if the selection is based on self-amplification or affinity “panning”. Bulk emulsification can be achieved using various laboratory techniques: with a stirring bar [32,35], using a tissue homogenizer (high-shear mixer) [36], by vortexing [37], and extrusion through a filter [38].

4.1. Compartment manipulation

Four basic on-chip unit operations carried out during microfluidic screening of biocatalysts are reagent mixing, compartmentalization, sometimes followed by droplet consolidation by gelation, incubation, and sorting.
Droplet formulation: Emulsions must be formulated to allow droplets to reliably maintain their contents during incubation and subsequent manipulation without cross contamination or sample loss [39]. Early research with mineral oil and silicon-based surfactant Abil EM 90 revealed transfer of fluorescent products into the continuous phase under certain conditions. However, the leakage of fluorophores can be reduced more than 10 times by adding 5% bovine serum albumin in the aqueous inner phase. Fluorinated oils such as Novec HFE7500 in combination with fluorinated surfactants offer an alternative carrier phase to prevent crosstalk [40].
On-chip mixing of inlet streams: On-chip mixing of genetic material (cells or TT mixtures) and the reagent is usually done immediately before compartmentalization, which allows the initiation of an enzymatic reaction just prior to droplet formation. On chip mixing involves mixing a suspension of cells or enzymes with a fluorogenic substrate and lysis agent using a mixing junction located upstream of the droplet generation junction. The mixing junction can have a T-, X- or Y- shape. In the case of a cross junction [41], cells suspended in a fermentation medium are injected through the main channel, while the substrate is delivered through two orthogonal side channels and the two streams are mixed by chaotic advection in the main channel [42]. In the case of a Y-junction [43], the cell suspension is delivered through one arm and the fluorogenic substrate through the second symmetrical arm of the Y-junction (Figure 5).
Droplet generation (compartmentalization): Droplet generation units for single (two-phase) emulsions can be based on T-junctions (cross-flowing streams), flow focusing junctions (elongated flow), co-flow streams, and step emulsification. The two crucial requirements in this step are that droplets are formed in dripping regime and a single-gene encapsulation is achieved, i.e. droplets must be “monoclonal” and monodisperse.
T-Junction: In a T-junction droplet generator, the continuous phase (CP) flows within the main channel and the dispersed phase (DP) is injected from a side channel intersecting at 90° (Figure 6 a). Droplets are generated due to shear forces induced by the continuous phase [44,45,46] or pressure force if the main channel is completely obstructed by the dispersed phase [47].
Co-flow droplet maker: Coflowing microfluidic droplet generators consist of two coaxial channels. The coaxial geometry is often formed by a smaller capillary tube being placed inside a larger concentric tube (Figure 6 b). The inner capillary supplies the DP whilst the outer tube delivers the CP fluid [48,49].
Flow-focusing junction: In this geometry, the DP flows in the middle channel and is enveloped by the CP coming from either side (Figure 6 c). The two liquid phases are usually forced through a small orifice downstream of the junction, and droplets are formed by viscous stress exerted on the inner phase by the surrounding outer phase flow. The geometry of these devices can vary from 3-D axisymmetric [50] to quasi-2D planar [51,52] to completely 2D planar. Axisymmetric devices provide a DP phase through a cylindrical inlet, which is hydrodynamically focused by the CP as both phases pass through a circular baffle. In all droplet generators mentioned above, the droplet size depends on fluid flow rates, channel geometry and stream compositions [52].
Microfluidic step emulsification: Microfluidic step droplet generators combine a shallow and narrow upstream microchannel delivering the DP and an abrupt (step-like) opening to a deep and wide reservoir (“well”) filled with the CP. The upstream narrow microchannel can terminate at the well or a wide terrace can be added between the narrow channel and the well to reduce the flow velocity of the DP and minimize their inertia [53]. Step droplet generators can easily be parallelized by connecting many shallow channels to the same well [54]. The well can contain a stagnant or perpendicularly flowing continuous phase [55]. In addition, narrow channels can be etched on the top surface of the substrate as micro-grooves [53] or through the entire cross section as straight-through channels [56]. In step emulsification, the droplet size in dripping regime is independent on the CP flow rate, since droplet instability is not achieved due to shear forces generated by the CP but due to imbalance of capillary pressure along the DP interface when the droplet is deformed at the step [57,58] (Figure 6 d).
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Figure 6. Lab on chip modules for microfluidic emulsifications (DP = dispersed phase, CP = continuous phase [59].
Figure 6. Lab on chip modules for microfluidic emulsifications (DP = dispersed phase, CP = continuous phase [59].
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The optimum droplet volume depends on the growth rate and incubation time of the cells. Using nanoliter- rather than picoliter-scale droplets is essential for screening of filamentous fungi [60]. Typically, the time from which Aspergillus spores germinate to the stage where they start to display detectable enzymatic activity is 24 h and optimum levels of activity are only reached after several days of incubation. The requirement for long incubation time combined with the rapid growth of the fungal hyphae makes screening filamentous fungi in picoliter droplets difficult because the hyphal tips exit 250 pl droplets in ~15 h. In contrast, encapsulating single spores in 18 nl droplets allows growth of the branched mycelial network for up to 24 h confined in the droplet with the hyphal tips exiting the droplets only after incubation for 32 h.
Droplet incubation: Droplet incubation can last from several minutes up to several weeks and can be achieved on-chip or off-chip (in a test tube) [43,61]. On-chip droplet incubation can be achieved using wavy channels (with an added benefit that droplet contents are mixed during incubation period), rectangular storage chambers [62] or microfluidic traps [63]. An array of pillars can be utilized along the side walls of the culture chamber (Figure 7 c) to allow carrier oil to flow while all droplets remain trapped inside, resulting in a highly packed droplet array inside the culture chamber [64].
After incubation, droplets are reinjected and sorted based on readout, which can be based on fluorescence, image analysis, light scattering, surface tension, electrochemical signal, and buoyancy.

4.2. Detection methods

Table 1. Detection methods used for sorting in microfluidics.
Table 1. Detection methods used for sorting in microfluidics.
Detection method Droplet size Sensitivity References
Absorbance High pL Mid µm [66]
Fluorescence intensity Low pL Low nM [67]
Raman spectroscopy Mid nL n.a. [68]
Electrochemistry Mid nL Low µm [69]
Mass spectrometry Mid nL Mid µm [70]
Light scattering High pL n.a. [71]
Surface tension High pL High nM [72]
Buoyancy Mid nL n.a. [73]

4.3. Sorting methods

Sorting includes signal detection and particle / droplet separation based on the intensity of target signal. The incubated droplets flow through the detection points one by one to identify the target signals, which is usually fluorescence intensity, but could be other physical property that can be linked with the enzyme activity [74]. The advantages of fluorescent activated droplet sorting (FADS) over FACS are the potential ability to screen for extracellular secretory products and the sorting of fungal spores at later stages of germination [75].
In dielectrophoretic droplet sorters droplets flow in carrier oil towards a Y-shaped junction. With no electric field, all drops flow into the waste channel, which offers lower hydrodynamic resistance than the second collect channel. To direct droplets into the collect channel, on chip electrodes are energized, creating an electrical field gradient, which generates a dielectrophoretic force (DEP) acting on the droplets (Figure 8).
Microfluidic cell sorters could be classified in two groups: active cell sorters that are based on an external power source (electric, acoustic, magnetic etc.) affecting the cells and forcing them to differentiate and passive cell sorters that rely on internal forces or geometrical patterns within the microchannels for separating cells [77].

5. Directed evolution of enzymes using microfluidics

5.1. Directed evolution of enzymes within microdroplets

5.1.1. Single emulsions

Whole cells or genetic material can be encapsulated in W/O droplets. Due to the external oil phase, W/O droplets are not compatible with commercially available FACS devices. To overcome this challenge, methods for on-chip droplet sorting have been implemented (Figure 9).
In the procedure shown in Figure 9, E. coli cells expressing the enzyme mutant library (red) are co-compartmentalized with the cell lysis solution and fluorogenic substrate. The cell encapsulation follows the Poisson distribution and single-cell compartmentalization can be achieved by adjusting the dilution of the cell-containing solution to the required level. There are several actively-controlled droplet production methods that present an alternative route to the production of droplets at similar rates and with the potential to improve the efficiency of single-cell encapsulation predicted by the Poisson distribution [80].
Upon droplet formation, the cells are lysed, and the enzyme reaction takes place within the droplet, leading to the accumulation of fluorescent product. Droplets exhibiting the highest fluorescence are sorted and plasmids are extracted and introduced into fresh E. coli cells to complete the directed evolution cycle. The developed protocol enables cell lysis in the droplets and allows the use of a wide variety of fluorescent substrates, regardless of their ability to penetrate the E. coli cell membrane. This feature is of particular importance, because most microtiter-based enzyme screening assays involve cell lysis prior to kinetic analysis.
Microfluidics can also be used for sorting in single emulsions of important industrial enzymes like glucose oxidase using a specially designed fluorescent assay that detects peroxide without bleaching. This was shown in research where glucose oxidase mutant with 2.1-fold higher activity towards glucose compared to wild-type enzyme was found [81]. A cellulase screening system based on fluorescence assay and droplet microfluidics in single emulsions was also developed on a similar microfluidic device [82].

5.1.2. Double emulsions

The process of in-vivo compartmentalization in bulk W/O emulsions and detection by FACS is described in Figure 10. (1) A gene library is transformed and cloned into E. coli, and (2) the encoded proteins are allowed to translate in the cytoplasm, or on the surface, of the bacterial cells. (3) Single cells are compartmentalized in the aqueous droplets of a W/O emulsion. (4) The fluorogenic substrate is added (through the oil phase), and the W/O/W emulsion is formed by emulsification of the primary W/O emulsion, enveloping the aqueous droplets with an intermediate layer of oil and providing an external aqueous phase. (5) Compartments containing the fluorescent product are sorted by FACS, and the cells embedded in them, together with the gene encoding the enzyme of interest, are isolated.

Microfluidic droplet sorting in double emulsions:

Droplet microfluidics can be used also for sorting in double emulsions. One research study shows development of the platform that based on microfluidics system with large double droplets that can encapsulate large mutant strains of filamentous fungi and obtaining strain with 2 fold higher α-amylase secretion capacity of A. niger (Figure 11) [75].
The role of Gel-MA is to prevent mycelial punctures and thus sustain prolonged culture. In another recent research a simple strategy that employs oil as a transient medium, water-water-oil double emulsions were generated by microfluidics using passive flow focusing and active pico-injection. Obtained droplets at high generation frequency of 2.4 KHz were used for cell encapsulation and fluorescence-activated droplet sorting [84].

5.1.3. Liposomes

Directed evolution of proteins can also be done using liposomes, lipid vesicle made from phospholipid bilayers. One of the early examples was investigation of in-vitro translation of green fluorescent protein within cell-sized giant unilamellar vesicles (GUV) by flow cytometry [85]. The authors proved that these liposomes provide ideal reaction environment that does not influence internal biochemical reactions (Figure 12).
GUVs suitable for directed evolution can be prepared in microfluidic devices using various methods. For example, there is report of using microfluidic-based mechanical droplet-splitting pipeline for the production of carrier-GUVs with diameters of ~2 μm that could encapsulate antibodies, or viruses [88]. In another microfluidic method GUVs with sizes from 10 to 30 µm that could encapsulate plasmids, smaller liposomes, mammalian cells and microspheres were produced using using PDMS-based lab-on-chips without surfactants [89]. GUVs can be also produced stepwise on a microfluidic assembly line [90], or via modified microfluidic method via water/octanol-lipid/water double emulsion droplets [91].

5.2. Directed evolution of enzymes within microgels

5.2.1. Microbeads

Another strategy enabling the use of FACS consists in generating droplets with a gellable polymer in which genes and either encoded enzymes or whole cells can be encapsulated. In this approach, after encapsulation of all reaction components in monodisperse droplets together with dissolved gel-forming polymers (agarose, alginate, etc.), the droplet contents solidify to microspheres made up of crosslinked hydrophilic polymers. A gel can be formed upon lowering the temperature, due to addition of oppositely charged small molecules (crosslinkers) or due to covalent interactions. Various crosslinking mechanisms used in microfluidic devices to form microgel particles are recently reviewed [92]. Polyelectrolytes of opposite charge can be deposited onto the charged surface of microgel particles (for example, based on charge interactions between negative alginate in the gel and positive polyammonium electrolyte) to create a size-selective shell which is permeable only for molecules with Mw < 2 kDa.
Weaver et al. [93] encapsulated mammalian, bacterial, and fungal single cells in FACS-compatible agarose beads with diameters of 20 to 90 μm by cooling liquid droplets [82]. After an incubation step in the growth medium and a staining step with fluorescent markers for biomass, the cell colonies were analyzed by FACS [94].
In this study, antibodies bound to the streptavidin-coated microbeads could immobilize the translated proteins. Upon translation, the emulsion droplets were ruptured to retrieve the microbeads and, subsequently, incubated with horseradish peroxidase (HRP) which bound to the proteins of interest via a ligand. In a second step, the beads were incubated with hydrogen peroxide and fluorescein tyramide, leading to the fluorescent labeling of the bead. FACS sorting of the microbeads enabled the identification of a protein with high affinity towards the ligand used in the screen [95].
That screening of active enzyme variants can be done within microbeads via their formation, or degradation was shown in the study where yeast cells with surface-displayed glucose oxidase were encapsulated within fluorescein labeled alginate microbeads formed by glucose oxidase activity and sorted by flow cytometry [96] . Similar approach was used amino acid oxidase coupled with horse radish peroxidase [97]. These results show that microbead gelation could be easily adapted for use in microfluidics and for a broad class of enzymes that can form [98] or degrade hydrogels.
One of the examples is evolution of phosphotriesterase using screening of gell-shell beads made by microfluidic device. Using this approach, more than 107 clones per hour from combinatorial libraries were screened, and a 20-fold faster mutant was isolated in less than one hour [99].

5.2.2. Microcapsules

Directed evolution can also be done in microcapsules produced by microfluidics. These microcapsules are used not only for directed evolution but also for controlled drug release, tissue engineering, and bacteria and cell encapsulation [100]. For example in recent research authors developed innovative high-throughput screening system for sortase F from P. acnes based on encapsulation of single cells and their lysis within Hollow-core polyelectrolyte-coated chitosan alginate microcapsules [101].

6. Directed evolution of biomaterials/biocomposites

One of the cornerstones of human technology has been using materials in our environment like wood, alginate, cellulose, silk, collagen etc. harvested from from living organism. Due to recent development in biotechnology and DNA synthesis and sequencing technologies we are able now to engineer these biomaterials for specific purpose we need. Using principles of directed evolution of enzymes it is possible now to perform experiments in rational engineering and directed evolution of protein materials [102]. For engineering biomaterials there are some recent reviews that partially cover use of microfluidics and directed evolution of biomaterials [103,104,105,106,107,108]. Some examples that use the principles of directed evolution and will be discussed here include evolution of protein containers from Lumazine Synthase from Aquifex aeolicus [109], de novo design of proteins forming polyhedral structures [110,111], design of block copolymers composed of silk protein domens [112], directed evolution of peptide-based material that can bind polylactic acid [113], and evolution of streptavidin-binding proteins for directed evolution of materials [114].

6.1. Directed evolution and de novo design of Protein Containers

In this research, authors performed four rounds of mutagenesis and selection in order to obtain a variant with a 10-fold higher loading capacity than the starting capsid for HIV protease [109]. They have used Lumazine synthase from Aquifix aeolicus, which forms icosahedral capsids consisting of 60 or 180 identical subunits. They did not use microfluidics, but this is one of the first examples of the directed evolution of biomaterials based on proteins. In another similar study rational design was used to make two-component 120-subunit icosahedral protein nanostructures similar to small viral capsids [111]. They have obtained structures using computational design and in vitro assembly of icosahedral complexes was confirmed by electron microscopy, small-angle x-ray scattering and x-ray crystallography. In a more recent study a novel 2.5 megadalton nucleocapsid was evolved using a deep mutational scanning library [110].

6.2. Protein engineering of material binding peptides

High-throughput screening is a valuable technique for the evolution of protein-based materials. It was shown in a recent study where streptavidin-binding proteins were evolved using selection-based screening technique for streptaviding binding in biofilms of elastin-like polypeptide block libraries [114]. Magnetic bead sorting was used for initial selection and biolayers coated with streptavidin within 96-shallow-well microtiter plates. In similar recent research, another example of directed evolution of peptide materials that can bind polylactic acid (PLA) was described [113]. Evolved material-specific binding protein (in this case with affinity for PLA) fused with polymer degrading enzymes can be generally applied for degradation of any specific material that is of high importance to environmental protection or directed evolution of polymer degrading enzymes.

6.3. Engineering of living materials (ELMs)

Advances in synthetic biology and materials science gave rise to a new form of materials named engineered living materials (ELMs). ELMs will have capabilities similar to natural living materials, like growth, self-organization, and self-repair, but with tailored functions. ELMs could be used for environmental bioremediation, as biomedical therapeutics, in situ biosensing etc. There are reviews covering existing ELMs and available synthetic biology tools [115,116]. There are three major categories of ELMs using cells and: proteins, non-ribosomally synthesized structural polymers, and biominerals [117]. Proteins for designing ELMs are spider silk, keratin, elastin and collagen. Desing of these ELMs is promising due to the developed directed evolution methods for proteins based on flow cytometry and microfluidics. Non-ribosomally synthesized structural polymers are polysaccharides (alginate, agarose, pectin, chitosan), polyesters, polyamides and inorganic polyanhydrides [118]. ELMs based on biomineralization can be made by bacteria producing inorganic materials in the form of precipitated minerals [119]. Also, it is possible to express specific peptides on the surface of cells that facilitate calcite precipitation [120].
Use of microfluidics for engineering of living materials is gaining more and more attention due to the precise control of fluids for complex micromanipulations that enables in situ encapsulation, incubation and manipulation of living cells within microcapsules or microbeads with highly uniform size [121]. Usually, cells are encapsulated with droplet microfluidics by mixing a hydrogel precursor with living cells as a dispersed phase and intersecting the precursor with a continuous phase in channels to produce droplets that later solidify into living hydrogel beads [122]. Polymers that are usually used are alginate, photo crosslinked hydrogels, agarose, gelatin, etc [103].

7. Conclusion

These research studies prove that protein engineering and directed evolution of enzymes can be performed using microfluidic lab on chip devices and recent modular microfluidics using various encapsulation methods in microdroplets, microbeads, and microcapsules using various detection methods like fluorescence, Raman spectroscopy, absorbance, mass spectroscopy and even the same beads and capsules as substrates where enzymes/cells are encapsulated. We have also shown that there are trials of using microfluidics for engineering novel protein-based and living materials that offer new possibilities to evolve novel materials, biocatalysts, and enzymes [102].

Author Contributions

Conceptualization, G.V. and R.P.; writing—original draft preparation, N.P.K., R.O., G.V. and R.P. writing—review and editing, G.V. and R.P.; visualization, N.P.K.; supervision, R.P.; funding acquisition, R.P. All authors have read and agreed to the published version of the manuscript.” Please turn to the for the term explanation. Authorship must be limited to those who have contributed substantially to the work reported.

Funding

This research was funded by the Ministry of Science, Technological Development and Innovation of the Republic of Serbia, Grant No. 451-03-47/2023-01/200168 (University of Belgrade-Faculty of Chemistry).

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

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Figure 1. Fabrication methods for fabrication pf microfluidic chips.
Figure 1. Fabrication methods for fabrication pf microfluidic chips.
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Figure 2. A Lego®-like microfluidic building block with the bottom surface (a) and the top surface (b). An example of a building block assembly is also shown. .
Figure 2. A Lego®-like microfluidic building block with the bottom surface (a) and the top surface (b). An example of a building block assembly is also shown. .
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Figure 3. Monodispersed W/O/W and O/W/O double emulsion droplets with controlled size and number of inner droplets generated by connecting two CNC-milled modules by a glass capillary [29].
Figure 3. Monodispersed W/O/W and O/W/O double emulsion droplets with controlled size and number of inner droplets generated by connecting two CNC-milled modules by a glass capillary [29].
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Figure 4. (a) A microchip with two miniaturized male luer fittings; (b) An H-shaped microchannel insert with two miniaturized female luer fittings; (c) Plugging the chip into two separate microchannel inserts. The male luer fitting was designed to plug into the female luer fitting with a diameter of 1 mm [30].
Figure 4. (a) A microchip with two miniaturized male luer fittings; (b) An H-shaped microchannel insert with two miniaturized female luer fittings; (c) Plugging the chip into two separate microchannel inserts. The male luer fitting was designed to plug into the female luer fitting with a diameter of 1 mm [30].
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Figure 5. Main strategies for on-chip mixing of inlet streams (cells and reagent) prior to droplet generation and cell screening.
Figure 5. Main strategies for on-chip mixing of inlet streams (cells and reagent) prior to droplet generation and cell screening.
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Figure 7. Strategies for on-chip droplet incubation: (a) wavy channel; (b) chamber; (c) wavy channel with pillars along the side wall to remove the continuous phase; (d) microwells for trapping droplets [65].
Figure 7. Strategies for on-chip droplet incubation: (a) wavy channel; (b) chamber; (c) wavy channel with pillars along the side wall to remove the continuous phase; (d) microwells for trapping droplets [65].
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Figure 8. Droplet sorting in dielectrophoretic microfluidic sorter based on fluorescence intensity [76].
Figure 8. Droplet sorting in dielectrophoretic microfluidic sorter based on fluorescence intensity [76].
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Figure 9. Microfluidic screening of cell lysate in W/O emulsions [78,79].
Figure 9. Microfluidic screening of cell lysate in W/O emulsions [78,79].
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Figure 10. High-throughput screening of enzyme libraries by Fluorescence-Activated Sorting of Single Cells in double emulsions [83].
Figure 10. High-throughput screening of enzyme libraries by Fluorescence-Activated Sorting of Single Cells in double emulsions [83].
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Figure 11. Core-shell droplet-based microfluidic screening system for filamentous fungi [75].
Figure 11. Core-shell droplet-based microfluidic screening system for filamentous fungi [75].
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Figure 12. Preparation of giant unilamellar vesicles by centrifugation of W/O emulsion placed on the top of an external aqueous phase. Monodispersed W/O emulsion can be produced by microfluidic emulsification [86,87].
Figure 12. Preparation of giant unilamellar vesicles by centrifugation of W/O emulsion placed on the top of an external aqueous phase. Monodispersed W/O emulsion can be produced by microfluidic emulsification [86,87].
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Copyright: This open access article is published under a Creative Commons CC BY 4.0 license, which permit the free download, distribution, and reuse, provided that the author and preprint are cited in any reuse.
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