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PAPER

Cite this: Lab Chip, 2018, 18, 323

Received 10th October 2017,Accepted 6th December 2017

DOI: 10.1039/c7lc01088e

rsc.li/loc

High-throughput double emulsion-basedmicrofluidic production of hydrogel microsphereswith tunable chemical functionalities towardbiomolecular conjugation†

Eric Y. Liu,a Sukwon Jung,a David A. Weitz, b

Hyunmin Yi*a and Chang-Hyung Choi *c

Chemically functional hydrogel microspheres hold significant potential in a range of applications including

biosensing, drug delivery, and tissue engineering due to their high degree of flexibility in imparting a range

of functions. In this work, we present a simple, efficient, and high-throughput capillary microfluidic ap-

proach for controlled fabrication of monodisperse and chemically functional hydrogel microspheres via

formation of double emulsion drops with an ultra-thin oil shell as a sacrificial template. This method utilizes

spontaneous dewetting of the oil phase upon polymerization and transfer into aqueous solution, resulting

in polyIJethylene glycol) (PEG)-based microspheres containing primary amines (chitosan, CS) or carboxylates

(acrylic acid, AA) for chemical functionality. Simple fluorescent labelling of the as-prepared microspheres

shows the presence of abundant, uniformly distributed and readily tunable functional groups throughout

the microspheres. Furthermore, we show the utility of chitosan's primary amine as an efficient conjugation

handle at physiological pH due to its low pKa by direct comparison with other primary amines. We also re-

port the utility of these microspheres in biomolecular conjugation using model fluorescent proteins, R-

phycoerythrin (R-PE) and green fluorescent protein (GFPuv), via tetrazine–trans-cyclooctene (Tz–TCO) liga-

tion for CS-PEG microspheres and carbodiimide chemistry for AA-PEG microspheres, respectively. The re-

sults show rapid coupling of R-PE with the microspheres' functional groups with minimal non-specific ad-

sorption. In-depth protein conjugation kinetics studies with our microspheres highlight the differences in

reaction and diffusion of R-PE with CS-PEG and AA-PEG microspheres. Finally, we demonstrate orthogonal

one-pot protein conjugation of R-PE and GFPuv with CS-PEG and AA-PEG microspheres via simple size-

based encoding. Combined, these results represent a significant advancement in the rapid and reliable fab-

rication of monodisperse and chemically functional hydrogel microspheres with tunable properties.

Introduction

Hydrogel microparticles hold significant potential in a broadrange of applications including biosensing,1,2 drug delivery3,4

and tissue engineering.5,6 This potential is gaining more trac-tion from recent advances in materials and fabricationmethods that have enabled the reliable production of micro-particles with controlled properties including shape, size,

chemical function, and porosity.7,8 For example, batchprocessing-based photolithographic9,10 and micromoldingtechniques11–13 allow for reliable fabrication of chemicallyfunctional microparticles with precise control over particleshape and dimensions. Meanwhile, microfluidic techniquesallow for the high-throughput fabrication of microparticleswith readily tunable size and complex geometries.14 Precisecontrol over microfluidic flows and inclusion of micro-RNA's,15 cells16 or other functional components inprepolymer solutions have enabled the development of highthroughput fabrication techniques for multiplexed microparti-cle arrays for biosensing or tissue engineering applications.

However, there still exist critical challenges and gaps indeveloping these potent microparticle systems despite recentadvancements. Notably, while there are numerous reports ofuniform microparticles imparted with magnetic, biodegrad-able, encoding, or DNA-, drug- or cell-carrying function,14

Lab Chip, 2018, 18, 323–334 | 323This journal is © The Royal Society of Chemistry 2018

aDepartment of Chemical and Biological Engineering, Tufts University, Medford,

Massachusetts, 02155, USA. E-mail: hyunmin.yi@tufts.edub John A. Paulson School of Engineering and Applied Sciences, Harvard University,

Cambridge, Massachusetts, 02138, USAc Division of Cosmetic Science and Technology, Daegu Haany University,

Gyeongsan, 38610, Republic of Korea. E-mail: cchoi@dhu.ac.kr

† Electronic supplementary information (ESI) available: Raw epifluorescencemicrographs of protein-conjugated chitosan microspheres in Fig. 6; negativecontrols for one pot assembly in Fig. 7. See DOI: 10.1039/c7lc01088e

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there has been a lack of reports of such microparticles withuniform chemical functionality.17 In addition, while provid-ing reliable routes to uniform and chemically functional hy-drogel microspheres, the batch-based nature of micro-molding techniques limits throughput.18 The incorporationof tunable and uniform chemical functionalities into hydro-gel microparticles in a rapid, high-throughput microfluidicfabrication method would thus represent a significant stepforward, for example by enabling facile post-fabrication bio-functionalization with molecular probes.

Our approach to addressing these challenges is an inte-grated fabrication–conjugation scheme utilizing a rapidmicrofluidic fabrication of chemically functional hydrogelmicrospheres, followed by efficient chemical reactions forbiomolecular conjugation. In this report, we first demon-strate fabrication of polyIJethylene glycol) (PEG)-based hydro-gel microspheres with two potent chemically functionalgroups, namely primary amines from an amino-polysaccharide chitosan (CS) and carboxylates from acrylicacid (AA), via a capillary microfluidic setup based on a novelsacrificial double emulsion-based approach (Fig. 1a). Unlikeconventional microfluidics-based methods, this microfluidicsetup produces double emulsion drops with an ultra-thin oilshell as templates, leading to reliable manufacturing ofhighly monodisperse hydrogel particles with minimal use ofoil phase and without harsh washing steps. Simple fluores-cent labelling reactions show that the primary amines of CSare efficient toward amine-reactive chemistries near physio-logical pH conditions due to its low pKa19 value by directcomparison with amines with typical pKa values found inbiomolecules. Next, we utilize a bright red fluorescent proteinR-phycoerythrin (R-PE) as a model protein to demonstratebiomolecular conjugation via a rapid and high yield bio-orthogonal reaction as well as the commonly enlistedcarbodiimide chemistry, and utilize these schemes to thor-oughly examine and compare the bioconjugation kineticsand polymer network structures of CS-PEG and AA-PEGmicrospheres. Finally, we show the potential for one-pot bio-molecular assembly using the two mutually orthogonal reac-tions with simple size-based encoding. Combined, we believethat the results in this report illustrate a significant step for-ward for programmable high-throughput fabrication and bio-molecular conjugation approaches that can be readily ex-panded to overcome limitations in a range of applicationareas including rapid biosensing, medical diagnostics andbiological threat detection in suspension array formats re-quiring minimal sample volume.

Results and discussionRapid capillary microfluidic fabrication of functionalhydrogel microspheres

We first demonstrate a potent and high-throughput capillarymicrofluidic-based fabrication of hydrogel microspheresusing a sacrificial double emulsion-based approach, asshown in Fig. 1a. The capillary device consists of three circu-

lar capillaries with different tapered orifice sizes inserted intoa square capillary.20 The injection and collection capillariesare chemically treated to impart hydrophobic and hydrophilicsurface properties, respectively. Thus, the oil and the aqueousphases preferably wet and can flow smoothly through eachcapillary. Both capillaries are precisely aligned and closely po-sitioned with the distal end of the collection capillary sealedto the square capillary. In addition, a small tapered circularcapillary is inserted into the injection capillary to supply thepolymerizable aqueous phase. Finally, the collection capillaryis connected to a polyethylene microtubing where ultraviolet(UV) light-induced free radical polymerization occurs beforethe polymerized hydrogel microspheres are collected in acontainer (Experimental).

The coaxial biphasic flow in the confined injection capil-lary results in a thin oil layer fully surrounding the

Fig. 1 One-step capillary microfluidic fabrication of chemicallyfunctional PEG microspheres via double emulsion drops with ultra-thinoil layer. (a) Schematic diagram of the glass capillary microfluidic de-vice used to prepare double emulsion drops containing PEGDA, photo-initiator, inert PEG600 porogen and either CS or AA in the inner-mostdrop. These drops are polymerized via UV light-induced free radicalpolymerization and collected in an aqueous wash to dewet the oillayer from the polymerized microspheres. (b) Brightfield micrograph ofthe formation of double emulsion drops. (c) Brightfield micrograph ofmicrospheres with uniform size. Scale bar represents 100 μm. (d) Sizedistribution of microspheres. (e) Plot of microsphere diameter vs. flowrate of the continuous phase showing control over microsphere size.

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polymerizable fluid due to strong affinity of the oil phase tothe hydrophobic inner wall of the injection capillary. An addi-tional aqueous continuous phase is injected through theinterstices between the square and the circular injection cap-illaries from the same side of the injection capillary, whilethe interstices of the square and circular collection capillariesat the distal end are sealed, preventing leakage flow and thusmaking strong shear force toward the collection capillary.The coaxial flow from the injection capillary is emulsified bythe continuous aqueous phase near the injection capillary'sexit, resulting in monodisperse double emulsion drops withthe ultra-thin oil layer flowing through the collection capil-lary. These drops are then exposed to UV irradiation (esti-mated exposure time: 5 s) in the end of polyethylene tubing(10 cm) that is connected with the collection capillary (5 cm),forming crosslinked polymeric hydrogel microspheres coatedwith the oil shell in a continuous manner without any block-age in the injection capillary from undesired UV exposure tothe polymerizable fluid. When these as-formed drops are dis-persed in the collection container containing deionized (DI)water, the oil shells start to dewet from the surfaces of thedrops, leaving hydrogel microspheres,16 while the separatedoil drops immediately migrate to the top due to buoyancy(ρhexadecane = 0.770 g mL−1 and ρ10% PEGDA aq. = 1.01 g mL−1).

As shown in the photomicrograph of Fig. 1b, this simpleyet potent setup yields exceedingly uniform polyIJethyleneglycol) (PEG) hydrogel microspheres in a high throughputmanner (i.e., approximately 400 spheres per s) as further il-lustrated in the brightfield micrograph of Fig. 1c. Detailedsize analysis of a representative batch of the as-preparedmicrospheres in Fig. 1d shows coefficient variation of lessthan 1.2%, clearly indicating the uniformity of the micro-spheres and robustness of the microfluidic setup. Further-more, the size of the microspheres is readily controlled bytuning simple parameters; for example, Fig. 1e shows thatone can create microspheres with highly uniform diametersin 65–103 μm ranges simply by changing the flow rate of theaqueous continuous phase (5–30 mL h−1).

Compared to conventional single emulsion (i.e. water-in-oil emulsion) based approaches,21,22 this approach enablesproduction of hydrogel microspheres without any extra wash-ing or separation procedure due to the aqueous continuousflow and spontaneous dewetting of the sacrificial oillayer.16,23 This thus eliminates the use of large amounts ofimmiscible oil phase as the carrier fluid, making the processsimple and cost-efficient.16 In short summary, the results inFig. 1 demonstrate a facile high-throughput fabrication tech-nique for readily controllable manufacturing of hydrogelmicrospheres.

Chemical functionality of chitosan-polyIJethylene glycol) (CS-PEG) microspheres

Next, we use fluorescent labelling to demonstrate the chemi-cal functionality and uniform incorporation of chitosan inthe PEG (CS-PEG) microspheres, as shown in Fig. 2. For this,

we fabricated CS-PEG microspheres by including short chainchitosan (Mn 5 kDa) in the PEGDA prepolymer mixture. Theas-prepared CS-PEG microspheres were exposed to anN-hydroxysuccinimidyl (NHS) ester form of fluorescein, asshown in the schematic diagram of Fig. 2a. The unsharedpair of electrons from CS's amine attacks the electron-deficient carbonyl group of the NHS-fluorescein, leading tothe formation of stable amide bond (i.e. amidation) in anacyl substitution reaction.24 This simple reaction thus is auseful tool to examine the presence, chemical functionalityand spatial distribution of CS in CS-PEG microspheres.

First, the brightfield micrographs in the first row ofFig. 2b show consistently uniform sizes of the CS-PEG micro-spheres with varying CS contents upon reaction with NHS-fluorescein, consistent with the results in Fig. 1. Next, theepifluorescence (top view) micrographs in the second row ofFig. 2b show minimal fluorescence for microspheres pre-pared without CS and increasingly bright and uniform

Fig. 2 Fluorescent labelling with CS-PEG microspheres. (a) Schematicdiagram of NHS-fluorescein labelling with CS-PEG microspheres. (b)Brightfield (top row) and epifluorescence (bottom row) micrographs offluorescently labelled CS-PEG microspheres prepared with 0–0.8 wt%CS. Scale bars represent 200 μm. (c) Plot of total fluorescence intensityvs. wt% chitosan in the prepolymer solution showing consistent incor-poration of chitosan in the microspheres.

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fluorescence for those prepared with increasing CS contentfrom 0.2–0.8%, indicating that there is minimal non-specificadsorption of fluorescein and that CS is consistently retainedwithin the microspheres. The fluorescence among micro-spheres in each condition appears uniform, further indicat-ing reliable fabrication of the microspheres and consistentincorporation of CS. Moreover, the fluorescence within eachmicrosphere appears uniform for each condition, suggestinguniform distribution of CS throughout each microsphere.Due to its small size (M.W. 473.4 Da), NHS-fluorescein rap-idly penetrates through the polymer networks and reacts withthe available primary amine moieties of CS, which are pre-sumably incorporated with the polymer networks via Michaeladdition in a stable manner.25 This result thus clearly illus-trates the chemical functionality and uniform incorporationof CS in the microspheres.

We then quantified the total fluorescence for CS-PEGmicrospheres prepared with varying CS contents, as shown inFig. 2c. For this, we used ImageJ software to measure the av-erage fluorescence intensity of a sphere multiplied by its as-sociated area to generate a total fluorescence intensity value,averaged over at least 10–20 spheres per condition examined.The total fluorescence intensity plot of Fig. 2c shows strongpositive correlation with higher CS content in the prepolymersolution. This result indicates that CS is consistently incorpo-rated in the microspheres, consistent with our recent studieson micromolding-based CS-PEG microparticles.24,26

In short summary, the simple fluorescein labelling resultsin Fig. 2 illustrate abundant primary amine functionality andconsistent, uniform incorporation of CS in CS-PEG micro-spheres fabricated via our rapid capillary microfluidicprocess.

Chemical functionality of acrylic acid-polyIJethylene glycol)(AA-PEG) microspheres

We next demonstrate carboxylate functionality and uniformincorporation of acrylic acid (AA) in AA-PEG microspheres viafluorescent labelling with EDC/NHS chemistry, as shown inFig. 3. For this, we fabricated AA-PEG microspheres underidentical fabrication conditions as the CS-PEG microspheres(Fig. 2), replacing CS with 0–0.5 vol% AA in the prepolymermixture. We then exposed the as-prepared AA-PEG micro-spheres to a 4 to 1 mixture of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) andN-hydroxysuccimidic acid (NHS), which converts the carboxyl-ates in the AA-PEG microspheres into NHS esters27 as shownin the schematic diagram of Fig. 3a. These NHS ester-activated microspheres were then reacted with a primaryamine-containing dye fluorescein glycine amide (FGA) viaacyl substitution reaction. Similar to the NHS-fluorescein re-action with the CS-PEG microspheres shown in Fig. 2 yet inreverse direction (i.e. amine group on the fluorescent markerinstead of CS), this widely used reaction scheme is thus use-ful in examining the presence and chemical functionality ofthe carboxyl groups in microspheres.

First as shown in the brightfield micrographs in the toprow of Fig. 3b, the FGA-labelled AA-PEG microspheres arehighly uniform in size, again indicating the robust nature ofthe capillary microfluidic process. Of note, the AA-PEG micro-spheres display intense yellow colour upon fluorescent label-ling via EDC/NHS reaction, suggesting abundant carboxylatefunctionality. Next, the epifluorescence micrographs (topview) in the bottom row of Fig. 3b show bright and uniformfluorescence among and within AA-PEG microspheres, indi-cating that AA-PEG microspheres contain abundant and uni-formly incorporated carboxylates. Meanwhile, AA-PEG micro-spheres without NHS ester activation and microsphereswithout AA show minimal fluorescence upon identical EDC/NHS and FGA exposure (middle and rightmost images at thebottom row of Fig. 3b respectively). These two results indicatethat EDC/NHS is required for the conversion of carboxylatesinto NHS esters for the covalent coupling with the primaryamine of FGA, and that there exists minimal non-specific ad-sorption of FGA with the PEG microspheres, consistent withour recent report on micromolding-based microspheres.18 Insummary, the simple fluorescein labelling results in Fig. 3clearly illustrate the abundant carboxylate functionality anduniform incorporation of AA in the AA-PEG microspheres.

Effect of pH: CS as an efficient conjugation handle

Upon examining the chemical functionality of our CS-PEGand AA-PEG microspheres, we next studied the effect of pHin the NHS ester–amine reaction on the microspheres' conju-gation efficiency. Briefly, we hypothesized that CS is a sub-stantially more efficient conjugation handle at lower and/orphysiological pH due to the difference in pKa of chitosan'sprimary amines (6.419) with other primary amines (i.e., FGA's

Fig. 3 Fluorescent labelling with AA-PEG microspheres. (a) Schematicdiagram of the EDC/NHS activation followed by FGA labelling with AA-PEG microspheres. (b) Brightfield (top row) and epifluorescence (bot-tom row) micrographs of FGA labelling and negative controls. Scalebars represent 200 μm.

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pKa 8.228). For this, we carried out the fluorescent labellingreactions described in the previous results (Fig. 2 and 3) atvarying pH (6–9) conditions. As shown in the epi-fluorescence micrographs in the top row of Fig. 4a, CS-PEGmicrospheres labelled with excess NHS-fluorescein displayuniform and increasingly bright fluorescence with increas-ing pH.

Similarly, in the bottom row of Fig. 4a, NHS ester-activated AA-PEG microspheres labelled with the identical ex-cess concentration of FGA also display uniform and increas-ingly bright fluorescence with increasing pH. These trends re-flect the increasing nucleophilic character with increasing pHof the primary amines within the CS-PEG microspheres orFGA, thereby increasing the reaction rate relative to that ofhydrolysis,29 and thus increasing the fluorescence intensityof the microspheres.

The normalized fluorescence intensity measurements inFig. 4b provide further semi-quantitative comparison of thisobservation. Here, the fluorescence intensity of CS-PEGmicrospheres significantly increases from pH 6–7, reaching70% maximum value at pH 7 and 80% at pH 8, while thefluorescence intensity of AA-PEG microspheres reaches only40% of maximum value at pH 7 and 70% at pH 8. This cleardifference in the conjugation efficiency at neutral pH (blackarrows) arises from the uniquely low pKa (6.4)19 of chitosan'sprimary amine compared to that of FGA (pKa 8.2).28 That is,a significant portion of primary amines in CS aredeprotonated at pH 7 compared to those of FGA, resulting inhigher fluorescence in CS-PEG microspheres. At pH 8, a sig-

nificant portion of the primary amines of the FGA moleculesbecome deprotonated, yielding higher fluorescence intensityof the AA-PEG microspheres. This result highlights the effi-ciency of chitosan's primary amines as conjugation handlesat lower pH compared to those of primary amines commonlypresent in biomacromolecules such as from N-termini (pKa7.7) and lysines (pKa 10.5) of proteins.30

Selective protein conjugation with CS-PEG and AA-PEGmicrospheres

To investigate the utility of the microspheres for bio-functionalization, we next examined protein conjugation withCS-PEG and AA-PEG microspheres using a large fluorescentmodel protein R-phycoerythrin (R-PE) via two conjugation re-action schemes. For the CS-PEG microspheres, we utilizedbiorthogonal tetrazine–trans-cyclooctene (Tz–TCO) chemis-try31,32 to covalently couple the primary amines of the micro-spheres with those of the R-PEs.

For this, we separately activated the primary amines in CS-PEG microspheres with Tz-PEG5-NHS ester and the primaryamines on R-PE with TCO-PEG4-NHS ester respectively, asshown in the schematic diagram of Fig. 5a. These Tz-activated CS-PEG microspheres were then exposed to theTCO-modified R-PE and imaged via epifluorescence

Fig. 4 Comparison of fluorescent labelling with CS-PEG and AA-PEGmicrospheres at pH 6–9. (a) Epifluorescence micrographs of fluores-cently labelled CS-PEG (top row) and AA-PEG (bottom row) micro-spheres at pH 6–9. Scale bars represent 200 μm. (b) Plot of normalizedtotal fluorescence intensity vs. pH for CS-PEG and AA-PEGmicrospheres.

Fig. 5 R-PE conjugation with CS-PEG and AA-PEG microspheres. (a)Schematic of Tz–TCO reaction for R-PE conjugation with CS-PEGmicrospheres. (b) Schematic of EDC/NHS reaction for R-PE conjuga-tion with AA-PEG microspheres. (c) Epifluorescence micrographs ofR-PE conjugation with CS-PEG (top row) and AA-PEG (bottom row)microspheres and negative controls. Scale bars represent 200 μm.

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microscopy. For AA-PEG microspheres, we utilized the EDC/NHS chemistry (Fig. 5b) to covalently couple the carboxylatesin the microspheres with the primary amines of the R-PEs.For both reaction schemes, we performed the conjugation re-actions for 8 h and at fixed R-PE concentration (2 μM).

As shown in the epifluorescence micrographs in thefirst column of Fig. 5c, both types of microspheres fluo-resce brightly and uniformly among microspheres uponR-PE conjugation, indicating that both types can readilybe coupled with large proteins (i.e. 240 kDa, 11 nm DH).

33

In addition, the uniformly high fluorescence within eachmicrosphere suggests that proteins have penetratedthroughout the polymer networks of the respective type ofmicrospheres, suggesting large hydrogel mesh size. Thefluorescence of the CS-PEG microspheres results from theinverse electron demand Diels–Alder reaction between theTz from the Tz-activated microspheres and the TCO onthe TCO-modified R-PEs,31,32 while the fluorescence of the

AA-PEG microspheres results from the acyl substitution ofNHS ester-activated AA-PEG microspheres with the primaryamines on R-PE.27,29 In direct contrast, microspheres showminimal fluorescence when reacted in the presence ofR-PE either without prior microsphere activation with Tzor requisite functional groups (second and third columnsof Fig. 5c). These results indicate that both CS-PEG andAA-PEG microspheres have minimal non-specific adsorp-tion with R-PE, presumably due to the non-fouling natureof the PEG, CS and AA.34,35 In turn, the minimal fluores-cence of these negative controls indicate that the fluores-cence observed in the first column of Fig. 5c is primarilydue to the covalent coupling of R-PE with the functionalgroups in CS-PEG and AA-PEG microspheres. The resultsin Fig. 5 thus demonstrate the utility of CS-PEG and AA-PEG microspheres in selective conjugation of large pro-teins via two conjugation reaction schemes, whilesuggesting macroporous nature of the polymer networks.

Fig. 6 Kinetic behaviour of R-PE conjugation with CS-PEG and AA-PEG microspheres. (a) Total fluorescence intensity vs. time plot and confocalmicrographs of R-PE conjugation with CS-PEG microspheres prepared with 0.1–0.8 wt% CS. (b) Total fluorescence intensity vs. time plot and con-focal micrographs of R-PE conjugation with AA-PEG microspheres prepared with 0.2–2 vol% AA.

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Protein conjugation kinetics

Motivated by the full penetration of R-PE proteins through bothtypes of microspheres observed in Fig. 5, we then thoroughlyexamined the kinetic and diffusive behaviours of CS-PEG andAA-PEG microspheres for the conjugation reactions with R-PEvia the Tz–TCO and EDC/NHS reaction schemes respectively, asshown in Fig. 6. Specifically, Tz-activated CS-PEG and NHSester-activated AA-PEG microspheres prepared with fixedPEGDA content and varying functional group contents wereconjugated with fixed concentration of TCO-modified R-PE andR-PE respectively for up to 4 h, with measurements taken atmultiple time points. Total fluorescence intensities from theresulting epifluorescence micrographs (ESI,† Fig. S1) were mea-sured at each time point, and confocal micrographs were alsotaken to examine the penetration depths of R-PE through themicrospheres' hydrogel networks over time.

First, as shown in the total fluorescence intensity vs. timeplot of Fig. 6a, the fluorescence intensities of CS-PEG micro-spheres prepared with 0.1–0.8 wt% CS rise rapidly, reaching60–70% of their respective maximum fluorescence intensitiesby 0.5 h. This rapid increase in fluorescence is most likelydue to the high reaction rate of the Tz–TCO reaction (820M−1 s−1),32 resulting in rapid coupling of TCO-modified R-PEwith the Tz groups within the CS-PEG microspheres. In turn,the rapid reaction rate of the Tz–TCO reaction enables TCO-modified R-PE to react with Tz sites even at very low concen-tration, making it possible to examine diffusion whileneglecting the effect of reaction rate (i.e., low Damköhlernumber).36 As expected, CS-PEG microspheres prepared withhigher CS content yielded higher initial rates of reaction andmaximum fluorescence intensities, due to the higher numberof functional groups (i.e. Tz-activated CS) in the micro-spheres. In addition, the fluorescence intensities of the 0.2,0.5, and 0.8% CS conditions continued to rise, onlyplateauing at 4 h, while the fluorescence intensity of the0.1% CS condition plateaued by 1 h. This suggests that theTz sites available within the microspheres prepared withhigher CS content were consumed by TCO-modified R-PEduring the 2–4 h period, while most of the Tz active sites inthe 0.1% CS-PEG microspheres seem to have been consumedby 1 h. Given the rapid rate of the Tz–TCO reaction, this ob-served “apparent” kinetic behaviour suggests that TCO-modified R-PEs are still diffusing into the centres of themicrospheres from 2–4 h for the 0.2–0.8% CS conditions,while the TCO-modified R-PEs have diffused throughout themicrospheres in the 0.1% CS condition by 1 h.

This differing kinetic behaviour observed via epi-fluorescence imaging is further confirmed by confocal micro-graphs on the right side of Fig. 6a. Specifically, the confocalmicrographs taken at the centres of CS-PEG microspheresprepared with 0.1–0.8% CS upon 0.5 h reaction show conju-gation of R-PE only near the outer regions for all the fourtypes of microspheres (first column). The centres of themicrospheres prepared with 0.1% CS display bright fluores-cence by 1 h and attain brighter, uniform fluorescence

throughout each microsphere by 2 h (bottom row), while cen-tres of microspheres prepared with higher CS content take in-creasingly longer time to display fluorescence. This indicatesthat R-PE can fully penetrate to the centres of microspheresprepared with 0.1% CS by 1–2 h. In contrast, the micro-spheres prepared with 0.2–0.8% CS show incomplete R-PEpenetration into their centres at 1 h (second column), withthe 0.2% CS case displaying deeper penetration than the0.5% or 0.8% CS cases. At 2 h, the 0.2–0.8% conditions allshow deeper R-PE penetration to the centres of the micro-spheres while maintaining a similar trend of 0.2% CSspheres displaying further penetration than 0.5% and 0.8%spheres (third column). Finally, by 4 h, all CS conditions dis-play bright and uniform fluorescence throughout the micro-spheres, indicating full penetration of R-PE to the centres ofthe microspheres. The slower penetration of R-PE into themicrospheres with higher CS content is likely due to a fewfactors. First, microspheres prepared with higher CS contenthave smaller mesh size, as observed in our recent reports.18

Specifically, more CS molecules in the prepolymer solutionshould lead to higher incorporation of CS molecules ascrosslinkers in the polymer networks via inefficient Michaeladdition reaction25 and hence smaller polymer mesh size.Second, microspheres prepared with higher CS content havemore active sites, allowing for more R-PEs to covalently bindto the outer regions first and sterically hinder the diffusionof subsequent R-PEs toward the centres of the microspheres.In essence, the kinetic behaviours observed in Fig. 6a isgoverned by the diffusion limitation of the R-PE within thepolymer networks of the CS-PEG microspheres. These resultsindicate that CS-PEG microspheres prepared with higher CScontent display higher protein binding capacity and reactmore rapidly, yet have smaller mesh size than the micro-spheres prepared with lower CS content.

Next, as shown in Fig. 6b, R-PE conjugation with AA-PEGmicrospheres via EDC/NHS chemistry displays different ki-netic and diffusive behaviours than CS-PEG microspheres.First, Fig. 6b shows that the fluorescence intensities of AA-PEG microspheres prepared with 0.2–2% AA increase slowlywith higher initial rates of reaction for the higher AA contentconditions. Specifically, the fluorescence intensities reach30–40% of maximum by 1 h for 0.2–2% AA conditions andplateau by 8 h. This is in direct contrast to the CS-PEG micro-spheres in Fig. 6a, which reach 70–80% of maximum fluores-cence by 1 h and plateau by 4 h. Furthermore, the AA-PEGmicrospheres displayed 5–10-fold lower maximum fluores-cence intensities compared to the CS-PEG microspheres.These results are likely due to the slower reaction rate of theacyl substitution reaction in contrast to the Tz–TCO reaction,along with competing hydrolysis and side reactions with theNHS ester groups in the AA-PEG microspheres consumingsome of the active sites.27,29

The diffusive behaviour of R-PE through the AA-PEGmicrospheres is also different from that of the CS-PEG micro-spheres, as further illustrated by the confocal micrographs tothe right of Fig. 6b. By 0.5 h, the centres of the microspheres

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prepared with 0.2–2% AA display uniform fluorescence, indi-cating that R-PEs have already diffused through the polymernetworks that are more macroporous than the CS-PEG micro-spheres (first column). The fluorescence of each type of AA-PEG microspheres continue to increase uniformly throughouteach microsphere over time (second and fourth columns),again in contrast to the CS-PEG microspheres. These resultsthus indicate that the R-PE conjugation with NHS ester-activated AA-PEG microspheres is more reaction-controlledthan the CS-PEG cases. We attribute this to larger AA-PEGmicrosphere mesh size arising from the poor polymerizationefficiency of AA and potential charge repulsion of the nega-tively charged carboxylates within the microspheres,37,38 inaddition to the slower and competing nature of the NHS es-ter–amine reaction.

In conclusion, the results in Fig. 6 highlight the differ-ences in kinetic and diffusive behaviour of protein conjuga-tion with diffusion governing the CS-PEG case and both dif-fusion and reaction governing the AA-PEG case.

Simultaneous one-pot protein conjugation

Finally, we demonstrate orthogonal one-pot conjugation oftwo model fluorescent proteins R-PE and GFPuv with CS-PEGand AA-PEG microspheres via the Tz–TCO and EDC/NHSchemistries and simple size-based encoding as shown inFig. 7. For this, we placed both 140 μm diameter CS-PEGmicrospheres and 200 μm diameter AA-PEG microspheres inone pot and added Tz-PEG5-NHS ester (reactive only to theprimary amines in the CS-PEG microspheres), as shown inthe schematic diagram of Fig. 7a. Upon washing excess Tz-PEG5-NHS ester, we then added EDC/NHS to the solution,which should react only with the carboxylates in the AA-PEGmicrospheres. Upon washing away excess EDC/NHS, we thensimultaneously added both GFPuv and TCO-modified R-PE ofvarying concentrations to the microsphere solution for 1 and4 h for the covalent coupling of TCO-modified R-PE with theCS-PEG microspheres and GFPuv with the AA-PEG micro-spheres respectively.

First, the brightfield micrographs in the first column ofFig. 7b show that the smaller CS-PEG and larger AA-PEGmicrospheres conjugated with 50 or 100 nM TCO-modifiedR-PE and 4 μM GFPuv are uniform and readily distinguish-able, illustrating simple size-based encoding. Next, thebrightfield-confocal overlays in the second column of Fig. 7bindicate the orthogonal conjugation of TCO-modified R-PEwith the CS-PEG microspheres and GFPuv with the AA-PEGmicrospheres. Meanwhile, the negative controls (ESI,† Fig.S2) show minimal cross-talk, providing further evidence ofthe orthogonal nature of the two reactions.

Finally, as shown in the confocal micrographs of the cen-tres of the microspheres in the third column of Fig. 7b, CS-PEG microspheres conjugated with higher TCO-modifiedR-PE content and longer time display increasing fluorescenceand R-PE penetration depth. Specifically, the CS-PEG micro-spheres for the 50 nM R-PE case (first row) show brighter

fluorescence on the outer regions than the centres upon 1 hreaction, while AA-PEG microspheres display uniform fluores-cence throughout at 1 h. At 4 h, both the CS-PEG and AA-PEGmicrospheres display brighter and completely uniform fluo-rescence, indicating complete diffusion of R-PE in CS-PEGmicrospheres and more proteins conjugated within bothtypes of microspheres. This trend is also observed in the CS-PEG and AA-PEG microspheres for the 100 nM R-PE condi-tion (third and fourth row), with CS-PEG microspheres show-ing incomplete R-PE penetration at 1 h and both types ofmicrospheres displaying brighter fluorescence and completepenetration of proteins by 4 h. As expected, at 1 h reaction

Fig. 7 One-pot conjugation of TCO-modified R-PE and GFPuv withCS-PEG and AA-PEG microspheres respectively. (a) Schematic diagramof one-pot conjugation. (b) Brightfield and confocal micrographs of 1and 4 h one-pot conjugation of 50 or 100 nM TCO-modified R-PE and4 μM GFPuv. Scale bars represent 200 μm.

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the higher 100 nM concentration of R-PE leads to thebrighter red fluorescence observed in CS-PEG microspherescompared to the 50 nM case above. These results indicate theorthogonal nature of the two reactions that enables simpleone-pot conjugation of two different proteins with respectivehydrogel microspheres.

Experimental

Hydrogen chloride, sodium hydroxide, Tween 20, 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide HCl (EDC),N-hydroxysuccinimide (NHS), 2-(4-morpholino)ethanesulfonicacid (MES), borate buffered saline (BBS) (20× concentrate, 50mM borate, pH 8.5), sodium phosphate monobasic anhydrous(99%), sodium phosphate dibasic anhydrous (99%), Luria-Bertani (LB) media, ampicillin, isopropyl β-D-1-thiogalactopy-ranoside (IPTG), and Bradford assay kits were purchased fromThermo Fisher Scientific (Waltham, MA). Bugbuster reagent andcentrifugal filter units (Amicon Ultra 0.5 mL) were purchasedfrom EMD Millipore (Billerica, MA). Hexadecane, polyIJethyleneglycol) (PEG, Mn 600 Da), polyIJethylene glycol) diacrylate(PEGDA, Mn 700 Da), chitosan oligosaccharide lactate (averageMn 5 kDa, >90% deacetylation), 2-hydroxy-2-methyl-propiophenone, acrylic acid, polyIJvinyl alcohol) (87–89% hydro-lyzed), n-octadecyltrimethoxyl silane, saline sodium citratebuffer (SSC) (20× concentrate, pH 7.0), and phosphate bufferedsaline (10 mM phosphate, 2.7 mM potassium chloride, 137 mMsodium chloride) pH 7.4 were purchased from Sigma-Aldrich(St. Louis, MO). 5- (and 6-) carboxyfluorescein succinimidyl ester(NHS-fluorescein) was purchased from Pierce Biotechnology(Rockford, IL). Dimethyl sulfoxide was purchased from ACROSOrganics (Geel, Belgium). R-Phycoerythrin (R-PE) was purchasedfrom Anaspec Incorporated (Fremont, CA). trans-Cyclooctene-polyIJethylene glycol)-N-hydroxysuccinimide ester (TCO-PEG4-NHS ester) and tetrazine-PEG5-NHS ester were purchased fromClick Chemistry Tools (Scottsdale, AZ). Fluoresceinyl glycine am-ide (FGA) was purchased from Setareh Biotech (Eugene, OR).Imidazole was purchased from Amresco (Solon, OH). ABIL EM90 was purchased from Evonik Industries (Germany).2-[MethoxyIJpolyethyleneoxy)propyl]trimethoxy silane was pur-chased from Gelest (Morrisville, PA). The glass capillaries werepurchased from AIT Glass (Rockaway, NJ). All chemicals wereanalytical grade and used without further purification.

Fabrication of capillary microfluidic device and dropgeneration

We first prepared injection capillaries by tapering circularglass capillaries (1B100-6, World Precision Instruments, Inc.,Sarasota, FL) with 560 μm inner diameter to 50 μm inner di-ameter using a micropipette puller (P-97, Sutter Instrument,Novato, CA). To render surface hydrophobicity, the innerwalls of the injection capillaries were treated withn-octadecyltrimethoxyl silane for 30 min and subsequentlywashed with ethanol. The circular injection capillary wasthen carefully inserted into the square capillary whose innerwidth (1.05 mm) is slightly larger than that of the outer diam-

eter of the injection capillary (1 mm). Next, the small taperedglass capillary with 10 μm inner diameter was prepared byheating and pulling a cylindrical capillary by hand using agas torch; this capillary was then inserted into the injectioncapillary for simultaneous injection of two immiscible fluids.Finally, the circular collection capillary was inserted into thesquare capillary from the other end; we also treated this col-lection capillary with 2-[methoxyIJpolyethyleneoxy)propyl]-trimethoxy silane to make the capillary wall hydrophilic. Dur-ing drop generation, the volumetric flow rate was controlledby syringe pumps (Harvard Apparatus, Holliston, MA) andthe production of the emulsion drops was observed using aninverted microscope equipped with a high-speed camera (Vi-sion Research Inc., Phantom V9.0, Wayne, NJ).

Fluorescent labelling with CS-PEG microspheres

As-prepared CS-PEG microspheres were reacted with 100 μMNHS-fluorescein in 5× saline sodium citrate (SSC) buffercontaining 0.05 vol% Tween 20 (SSC-TW20 buffer solution)(pH 7) or 20 mM sodium phosphate buffer (SPB) containing0.05 vol% Tween 20 (SPB-TW20) (pH 6, 7, 8, or 9) for 1 h,then washed with DI water (1×), DMSO (2×), and SSC-TW20buffer (pH 7) (3×) to remove excess unreacted and non-specifically bound NHS-fluorescein.

Protein modification with TCO

To prepare TCO-modified R-PEs, we first exchanged thebuffer solution of the R-PE with BBS (50 mM borate, 300 mMsodium chloride, pH 8.5) via centrifugal filtration units at 4°C. The R-PEs were then reacted with 20- or 50-fold molar ex-cess TCO-PEG4-NHS ester for 30 min at room temperatureand then purified from the unreacted excess TCO moleculesvia centrifugal filtration with phosphate buffered saline pH7.4. The final concentrations of the purified TCO-modifiedR-PE were measured using UV-visible spectroscopy (Evolution300 UV-vis spectrophotometer, Thermo Scientific) with thecharacteristic absorbance peak (565 nm) and molar extinc-tion coefficient (1.96 × 106 M−1 cm−1) of the R-PE.39

R-PE conjugation with CS-PEG microspheres

R-PE conjugation with CS-PEG microspheres was performedusing tetrazine–trans-cyclooctene chemistry.31 First, CS-PEGmicrospheres were activated with 500 μM tetrazine-PEG5-NHSester for 1 h in SSC-TW20 buffer (pH 7) at room temperature.The Tz-activated CS-PEG microspheres were washed withSSC-TW20 buffer (pH 7) (5×) and then reacted with 50 nM to2 μM TCO-modified R-PE for up to 8 h in SSC-TW20 buffer(pH 7) at room temperature, with samples collected at vari-ous time points. The microspheres were then washed withSSC-TW20 buffer (pH 7) (5–6×).

Fluorescent labelling with AA-PEG microspheres

To label the AA-PEG microspheres with FGA, we first acti-vated the AA-PEG microspheres with 0.4 M EDC and 0.1 M

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NHS in 20 mM MES buffer containing 0.05 vol% Tween 20(MES-TW20) (pH 6) for 15 min to provide a high number ofNHS ester functional groups as indicated in our recent re-port,18 and then washed away the unreacted excess withMES-TW20 buffer (pH 6) (2×) and SPB-TW20 buffer (pH 6, 7,8, or 9) (2×). Upon reaction with FGA in SPB (pH 6, 7, 8, or 9)for 1 h, the microspheres were then washed with DI water(1×), DMSO (2×) and SSC-TW20 buffer (pH 7) (3×) to removeexcess unreacted and non-specifically bound FGA.

Production and purification of green fluorescent proteins(GFPuv)

Green fluorescent protein modified for maximum fluores-cence under UV light (GFPuv) was generously provided by Dr.Chen-Yu Tsao and Dr. William E. Bentley at University ofMaryland in E. coli BL 21 harbouring the plasmid ptrcHisB::gfpuv,40 and obtained via standard protein expressionmethods. Briefly, the E. coli cells were cultivated in LB mediain flask cultures, and the GFPuv expression was induced byadding 0.4 mM IPTG. Upon harvesting and disrupting thecells with Bugbuster, a standard 1 mL immobilized metal af-finity chromatography (IMAC) column (GE Healthcare, Chi-cago, IL) was utilized to purify the hexahistidine-taggedGFPuv using 0.5 M imidazole as the displacer. The as-prepared GFPuv was quantified by a standard Bradford as-say41 and used in the protein conjugation studies withoutfurther purification.

R-PE conjugation with AA-PEG microspheres

To conjugate R-PE with the AA-PEG microspheres, we first ac-tivated the microspheres with 0.4 M EDC and 0.1 M NHS inMES-TW20 buffer (pH 6) for 15 min. Upon washing withMES-TW20 buffer (pH 6) (2×) and SSC-TW20 buffer (pH 8)(2×), we then reacted the NHS ester-activated AA-PEG micro-spheres with 2 μM R-PE for up to 8 h in SSC-TW20 buffer(pH 8) at room temperature, taking sample measurements atvarious time points. The microspheres were then washedwith SSC-TW20 buffer (pH 7) (5–6×).

One-pot GFPuv and TCO-modified R-PE conjugation with CS-PEG and AA-PEG microspheres

CS-PEG (140 μm diameter) and AA-PEG (200 μm diameter)microspheres were placed together in SSC-TW20 buffer (pH7). We then activated the spheres with 500 μM Tz for 1 h atroom temperature. Upon washing away excess unreacted Tzwith SSC-TW20 buffer (pH 7) (2×) and MES-TW20 buffer (pH6) (2×), we then activated the spheres with 0.4 M EDC and 0.1M NHS in MES-TW20 buffer for 15 min. We then washedaway excess with SSC-TW20 buffer (pH 8) and added 4 μMGFPuv and 50–100 nM TCO-modified R-PE to the solution for1 or 4 h. The GFPuv-conjugated AA-PEG and TCO-modified R-PE-conjugated CS-PEG microspheres were then washed withSSC-TW20 buffer (pH 7) (5–6×).

Image analysis

The fluorescently labelled and protein-conjugated micro-spheres were imaged with an epifluorescence microscope(Olympus BX51 equipped with a DP70 microscope digitalcamera, Centre Valley, PA) and a confocal microscope (LeicaDMIRE2 equipped with a TCS SP2 scanner, Wetzlar, Ger-many), all in SSC-TW20 buffer (pH 7). Epifluorescence micro-graphs of these microspheres were obtained with a 10× objec-tive lens under standard green (U-N31001), red (U-N31002),and UV (11000v3) filter sets (Chroma Technology Corp., Rock-ingham, VT). Confocal micrographs of the protein-conjugatedmicrospheres were obtained with a 10× and 20× objectivelens under 488 and 543 nm excitation with depth scan incre-ments of 10–20 μm. Fluorescence intensity and microspherediameter measurements were done using ImageJ image anal-ysis software.42

Conclusions

In this work, we presented a reliable and rapid one-stepmicrofluidic approach utilizing double emulsion drops withan ultra-thin sacrificial oil shell to fabricate highly monodis-perse hydrogel microspheres with tunable size and chemicalfunctionality in a simple, consistent and cost-efficient man-ner. Simple fluorescent labelling studies demonstrated theabundant and uniformly distributed primary amines (CS-PEG) or carboxylates (AA-PEG), with the primary amines ofCS representing an efficient conjugation handle at neutralpH. The utility of these microspheres for biomolecular conju-gation was demonstrated using large fluorescent proteinR-PE via Tz–TCO and EDC/NHS schemes, where confocalmicroscopy illustrated that diffusion primarily governs theprotein conjugation of CS-PEG microspheres with smallermesh size, while diffusion and reaction both govern the pro-tein conjugation of AA-PEG microspheres with larger meshsize. Finally, the one-pot protein conjugation results showedorthogonality and simple size-based encoding afforded byour integrated approach combining potent capillary micro-fluidic fabrication and selective bioconjugation. These resultsrepresent a significant advancement in the rapid and tunablefabrication of hydrogel microspheres with precise controlover size and chemical functionality over previous works, aswell as their potential for biomolecular applications throughthe rapid and selective conjugation of large proteins.

While not examined in this report, our fabrication–conju-gation approaches are highly modular and can be readily ex-tended to impart numerous other features. On the fabricationside, the choice of a range of monomers and multiple func-tional groups can impart additional functionality onto themicrospheres, such as N-isopropylacrylamide for thermal re-sponsiveness,43 higher acrylic acid content for pH-based re-sponsiveness,44 or caprolactone for biodegradability,45 toname a few. These functions could also be further enhancedthrough the inclusion of multiple compartments within themicrospheres for isolation of functions and other fea-tures.17,46 On the conjugation side, conjugation of various

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molecules of interest such as antibodies47 or peptideprobes48 with our chemically functional microspheres can en-able the capture of biospecific targets toward rapid bio-sensing or medical diagnostic applications. The chemicallyfunctional microspheres can also take advantage of a suite ofclick chemistries49 for the rapid and orthogonal conjugationof multiple probes or molecules of interest. We thus envisionthat our integrated fabrication–conjugation approach couldbe readily adopted to manufacture hydrogel microparticleswith multifaceted dimensional, functional and specific fea-tures for a wide range of applications.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

We gratefully acknowledge partial financial support by theNational Research Foundation of Korea (NRF) grant fundedby the Korea government (NRF-2017R1C1B2006237), the U.S.National Science Foundation (Grant # CBET-1703549), the Na-tional Science Foundation (DMR-1708729), and the HarvardMaterials Research Science and Engineering Center (NSFDMR-1420570), and the National Institutes of Health(R01EB014703).

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