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Microbial activation of Bacillus subtilis-immobilized microgel particles for enhanced oil recovery Han AM Son, Sang Koo Choi, Eun Sook Jeong, Bohyun Kim, Hyun Tae Kim, Wonmo Sung, and Jin Woong Kim Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.6b02010 • Publication Date (Web): 09 Aug 2016 Downloaded from http://pubs.acs.org on August 15, 2016

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Microbial activation of Bacillus subtilis-immobilized microgel

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particles for enhanced oil recovery

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Han Am Son,†,‡ Sang Koo Choi,§ Eun Sook Jeong,§ Bohyun Kim,§ Hyun Tae Kim,†,*

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Won Mo Sung,‡ Jin Woong Kim §,∥,*

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†

Korea Institute of Geoscience and Mineral Resources, Daejeon 34132, Republic of Korea

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‡

Department of Natural Resources and Environmental Engineering, Hanyang University,

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Seoul 04763, Republic of Korea

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§

Department of Bionano Technology, Hanyang University, Ansan 15588, Republic of Korea

∥

Department of Applied Chemistry, Hanyang University, Ansan 15588, Republic of Korea

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ABSTRACT: Microbially enhanced oil recovery involves the use of microorganisms to

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extract oil remaining in reservoirs. Here, we report fabrication of microgel particles with

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immobilized Bacillus subtilis for application to microbially enhanced oil recovery. Using B.

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subtilis isolated from oil-contaminated soils in Myanmar, we evaluated this microbe’s ability

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to reduce the interfacial tension at the oil–water interface via production of biosurfactant

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molecules, which eventually yield excellent emulsification across a broad range of the

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medium pH and ionic strength. To safely deliver B. subtilis into a permeable porous medium,

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in this study, these bacteria were physically immobilized in a hydrogel mesh of microgel

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particles. In a core flooding experiment, in which the microgel particles were injected into a

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column packed with silica beads, we found that these particles significantly increased oil

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recovery in a concentration-dependent manner. This result shows that a mesh of microgel

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particles encapsulating biosurfactant-producing microorganisms holds promise for recovery

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of oil from porous media.

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Keywords: microbially enhanced oil recovery, microgel particles, biosurfactants

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ACS Paragon Plus Environment

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1. INTRODUCTION

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Microbes have been used as an injected agent to enhance oil recovery from petroleum

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reservoirs because microbes are environmentally friendly and viable under harsh reservoir

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conditions, such as high temperatures and salinity.1–3 Some of them can produce surface-

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active compounds, which are usually called “biosurfactants.” These compounds lower the

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oil–water interface tension, thus facilitating the release of the remaining oil from rock

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pores.4–6 In terms of enhancement of oil recovery, utilization of microbes is indeed

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advantageous. Nevertheless, direct injection often causes accumulation of the microbes on

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the inner wall of the nozzle before arrival at the reservoir. If there is any microbial growth,

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then the injection nozzle can be completely clogged; this situation eventually hampers further

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injection of other materials into the reservoir.7,8 To get rid of the microbial plug, application

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of unsafe and undesirable high pressure to the nozzle is required. As a consequence, for

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practically useful microbially enhanced oil recovery, it is important to develop a

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straightforward approach that allows for efficient transport of the microbes into the oil

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reservoir.9

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To transport living species, such as microbes, to the target area, they first should be

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reliably protected from the harsh environmental factors. As a practical solution to this issue,

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the use of the microencapsulation technology has been widely evaluated in a variety of fields,

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including pharmaceuticals,10 biochemical sensors,11,12 cosmetics,13 and chemical catalyst

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reactions.14,15 Recent advances in the microencapsulation technology have been made by

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means of the microfluidic method.16 This protocol enables encapsulation of living species,

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such as living cells and microorganisms, in a mesh of microgel particles with high

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encapsulation efficiency without any harmful chemicals.17–19 The living species physically

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immobilized in a biocompatible hydrogel phase can adsorb and release biocompounds

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through the hydrogel mesh. This approach allows researchers to design a practical

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microcapsule system in which microbes are immobilized in the hydrogel phase while

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producing biosurfactants through the hydrogel mesh. The biosurfactants are expected to be

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readily released through the hydrogel mesh because their molecular size is smaller than the

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size of the mesh pores. This approach has advantages over conventional encapsulation

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methods because it does not require any specific stimuli to trigger the release of microbes

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from the capsule.20–22 2


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The ultimate goals of this study were to fabricate microgel particles with immobilized

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microbes by the microfluidic method and to determine whether they can be useful for

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microbial enhancement of oil recovery as a recovery agent. The microfluidic method allowed

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us to produce microgel particles with excellent size monodispersity. The particle size is

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determined by the shear forces and is adjustable by simply regulating the microfluidic device

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geometry and flow rate conditions. The microgel particles contain a Bacillus strain in their

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hydrogel phase. The Bacillus strain was collected from an oil reservoir in Myanmar. To

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experimentally demonstrate that the B. subtilis strain selected in this study produces

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biosurfactants, we evaluated the interfacial activity of B. subtilis strain at different pH and

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salinity levels. Finally, we carried out an oil recovery experiment using the microgel particles

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with B. subtilis immobilized in a porous medium. The microgel particles were injected into a

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column packed with silica beads filled with n-decane and their oil recovery capacity was

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evaluated quantitatively.

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2. EXPERIMENTAL SECTION

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2.1. Materials. Poly(ethylene glycol) diacrylate (PEGDA, Mw = 700 g/mol), sodium 4-

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vinylbenzenesulfonate

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poly(diallyldimethylammonium chloride) (average Mw = 20,000–350,000 g/mol, 20 wt% in

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H2O), poly(sodium 4-styrenesulfonate) (average Mw = ~70,000 g/mol), glycerin, and paraffin

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oil were purchased from Sigma-Aldrich (USA) and used without further purification. Cetyl

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PEG/PPG-10/1 dimethicone (Abil EM 90, Evonik, Germany) was used as received.

(NaSS),

2-hydroxy-2-methylpropiophenone

(Darocur

1173),

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2.2. Extraction of B. subtilis and incubation for biosurfactant production. The candidate

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organism for microbially enhanced oil recovery was isolated from the oil-contaminated soils

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collected in the Shwezetaw Formation (depth 460–550 m) in the Salin fore-arc basin,

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Myanmar. The microorganism, which grew on the minimal salt medium (MSM) containing a

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carbon source (glucose, 2 wt%) and a trace element solution (TES; 0.2 wt%), was isolated

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and identified as a strain of B. subtilis by means of 16S ribosomal RNA sequencing. Using

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the microorganism, we produced biosurfactant. The Erlenmeyer flasks containing MSM broth

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was sterilized by autoclaving at 120 °C for 20 minutes. Next, B. subtilis strain was

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incorporated and carbon source (glucose, 2 wt%) was added into the medium. The culture 3


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was incubated in a shaker incubator with a rotation speed of 120 rpm for 96 h at 34°C. In

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order to experimentally confirm the effective production of biosurfactants in incubated stock

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solution, the interfacial tension was measured with varying the concentration of B. subtilis in

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the stock solution at pH 8.0. Additionally, to evaluate whether the bacterial species really

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made a difference, Escherichia coli as a non-oil-tolerant bacterial species was incubated with

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the carbon source. Then, we investigated the changes of interfacial tension in the presence of

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E. coli under the same conditions as the case of using B. subtilis.

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2.3. Determination of interfacial tension between oil and water. The interfacial tension

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was measured to characterize the B. subtilis’s behavior at the oil–water interface at pH 4–10

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without NaCl and at pH 8.0 with 0–10 wt% NaCl. The interfacial-tension measurements were

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done with an optical tensionmeter (Biolin Scientific, Sweden) by reversed pendant drop

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shape analysis at room temperature. Incubated B. subtilis (0.9 g/L) was present in the aqueous

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phase at different pH levels and NaCl concentrations in each transparent rectangular cell.

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Then, a hooked needle was immersed in the aqueous phase in a rectangular cell, and the n-

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decane drop was hanging on the hooked needle. The shape of the n-decane drop is

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determined by means of the balance of forces including the interfacial tension between n-

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decane and the aqueous phase. The interfacial tension can be calculated by means of the

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Young-Laplace equation in software (One Attension, Biolin Scientific).

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2.4. Production of n-decane emulsions. Before emulsification test, B. subtilis was incubated

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with carbon source for production of biosurfactant. An n-decane emulsion was prepared with

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incubated aqueous solution containing B. subtilis (0.9 g/L). We controlled pH of the solution

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at 4 or 8, while changing the salinity by adding NaCl up to 10 wt%. To induce rapid phase

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separation, the volume of n-decane was set to 50% of the aqueous phase volume. Then, the B.

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subtilis suspension and n-decane were mixed for 2 min at 3000 rpm with an ultrasonic

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homogenizer. After sonication, phase separation of the emulsion samples was observed

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during storage at room temperature

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2.5. Fabrication of microcapillary microfluidic devices. To fabricate the devices, a

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cylindrical glass capillary (outer diameter 1.0 mm, World Precision Instruments, USA) was

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heated and pulled using a pipette puller (Model P-97, Sutter Instruments, USA). The thin tip 4


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of the tapered glass capillary was cut to the desired diameter using an MF 830 Micro Forge

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(Narishige, Japan). To make the surface of the round capillary tubes hydrophobic, 1 wt%

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hexyl-trimethoxysilane in toluene was circulated through the glass capillaries. Then, a

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tapered cylindrical glass capillary was inserted into a square capillary (inner diameter 1.0

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mm, Atlantic International Technology, USA). Each end of the square capillary was

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connected to a needle and pasted with an epoxy resin. The needle was connected to a

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polyethylene tube (PE-5, outer diameter 1.32 mm, Scientific Commodities, USA), which was

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again connected to a glass syringe (Hamilton Gastight, USA).

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2.6. Microfluidic synthesis of microgel particles. The dispersion fluid and outer fluid were

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injected through the interstices between the round injection capillary and the square

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collection capillary. The flow rates were precisely adjusted using the syringe pumps (Pump

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11 Elite, Harvard Apparatus, USA). The dispersion fluid was an aqueous monomer solution

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containing the incubated B. subtilis suspension (48 wt%), PEGDA (20 wt%), NaSS (10 wt%),

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glycerin (20 wt%), and Darocur 1173 (a photoinitiator, 2 wt%). The outer fluid consisted of

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paraffin oil and 0.5 wt% Abil EM 90. Formation of emulsion drops was monitored under an

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inverted microscope equipped with a high-speed camera (Phantom Miro eX2, Vision

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Research Inc., USA). Then, the emulsion drops were solidified by photo-polymerization

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under ultraviolet (UV) at a wavelength of 365 nm for 1 min (UV light, A&D Co., Korea), and

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the emulsion drops were transferred to microgel particles. The paraffin oil, remnant

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monomers, and other additives were thoroughly removed with isopropanol by repeated

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centrifugation at 5000 rpm. Finally, the microgel particles with immobilized B. subtilis could

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be recovered from the continuous phase of paraffin oil without the use of chemical solvents.

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2.7. Apparatus for core flooding experiment. We conducted the flooding experiment to

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quantitatively evaluate the microbial activity of the microgel particles with immobilized B.

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subtilis as a recovery agent. The outline of the flooding process is shown in Fig. 1. The

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apparatus for the core flooding experiment was fabricated by assembling an injection pump,

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an accumulator, a core holder, and a measuring cylinder. The piston plate was installed in the

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cylindrical accumulator, and the dispersion of the microgel particles filled the container

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located above the piston of the accumulator. When water was injected into bottom of the

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cylindrical accumulator by injection pump, the piston plate was pushed up. Then, the 5


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dispersion of the microgel particles was flowed into a column filled with silica glass beads 1

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mm in diameter. The fluid that flowed through the interstice of silica beads was collected in a

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volume-readable cylinder. Oil reservoir contained oil and water in the rock pore. Before the

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flooding experiment, water was injected into the silica beads, and then n-decane was injected

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into the water-saturated medium until no more water drained off. Then, the initial volume

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fraction of water and oil in the silica beads was calculated from the volume of water

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collected.

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3. RESULTS AND DISCUSSION

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B. subtilis is known as a bacterium that produces biosurfactants. A good example is

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surfactin.23 Analogous to typical surfactants, those biosurfactants have an amphiphilic

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molecular structure, so that they can assemble at the interface of fluids of different polarity,

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such as water and oil phases. There have been debates on how they manifest the surface

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activities. For instance, the biosurfactants may increase bioavailability of a hydrophobic

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substrate by increasing their apparent solubility in hydrocarbon compounds, thus easily

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desorbing them from the surface. Aside from that, they can regulate adhesion of

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microorganisms to a surface.24 Studies have shown that B. subtilis biosurfactants enhance

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aqueous solubility of hydrophobic substances; this change reduces the interfacial tension

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between oil and water. To elucidate in detail how the presence of B. subtilis biosurfactants in

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the aqueous phase affects the surface activity, in this study, we tried to directly measure

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changes in the interfacial tension between n-decane and water in various solution

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environments. Before the measurement of interfacial tension, B. subtilis solution was

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incubated for production of biosurfactant. To identify the production of biosurfactant in the

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B. subtilis solution, the interfacial tension was measured with varying dilution of B. Subtilis

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solution at pH 8 (Fig. 2).

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Here, the initial concentration of of B. Subtilis in solution before dilution was 0.9 g/L, and

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the declined concentration of B. Subtilis means the decreasing concentration of biosurfactant.

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As the aqueous phase is concentrated with B. subtilis, the interfacial tension sharply dropped

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from 49 dyne/cm to 18 dyne/cm. However, in the case of using the non-oil-tolerant bacterial

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species, E. coli, the interfacial tension retained unchanged irrespective of the concentration of

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E. coli, as shown in Fig. 2. This means that biosurfactant was effectively produced by the 6


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activity of B. subtilis. Also, we analysed the effect of pH and salinity on the B. subtilis

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biosurfactants. The result is shown in Fig.3. The interfacial tension between n-decane and the

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aqueous solution containing B. subtilis (0.9 g/L) decreased as the pH of the aqueous solution

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increased (Fig. 3A).The interfacial tension decreased from 32 to 18 dyne/cm as pH increased

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from 4 to 8. Across the pH range from 8 to 12, little or no change in the interfacial tension

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was observed. This is because the biosurfactants produced by B. subtilis strain have a

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tendency toward precipitation at low pH due to their low solubility under highly acidic

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conditions.25-27 Therefore, improved interfacial activity may be attained under neutral and

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alkaline conditions. The effect of salinity on the interfacial activity of B. subtilis was also

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evaluated by addition of NaCl (Fig. 3B). The interfacial tension gradually increased from 18

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to 23 dyne/cm as the concentration of NaCl reached 10 wt%. Even though there was some

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deterioration in the interfacial tension, this was an inspiring result because B. subtilis

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biosurfactants still showed interfacial activity even under such conditions of harsh salinity.

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Having confirmed that the presence of B. subtilis in the aqueous phase lowered the

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interfacial tension between oil and water by the formation of biosurfactants, we decided to

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incorporate B. subtilis solution, including produced biosurfactants, into an emulsification

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system. As shown in Fig. 4, the emulsion that was produced in the absence of solution

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containing B. subtilis (pure water) was easily separated into upper and lower phases (Sample

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1). The emulsions that were produced at pH 4 were also readily broken up within minutes

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(Samples 2–5). Even after 7-day storage, no improvement in the emulsion stability could be

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achieved. This finding implies that biosurfactants produced by B. subtilis couldn’t retain their

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own interfacial activity, because they tend to aggregate at pH 4. In contrast to the cases at low

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pH, when the emulsions were produced at pH 8, the emulsion stability was significantly

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enhanced (Samples 6–9). Even after long-term storage, the emulsion drops remained

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unchanged. This result indicates that B. subtilis biosurfactants have excellent interfacial

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activity at pH 8, in agreement with the results in Fig.3A. To assess the effect of salinity, we

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added various concentrations of NaCl to the aqueous solution containing B. subtilis. The

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emulsion index, i.e., the height of the emulsified layer as a percentage of the total height of

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the liquid column,28 was determined for ~20% to 5 wt% NaCl. The emulsion index

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remarkably increased to approximately 55% at a high NaCl concentration (10 wt% NaCl,

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Sample 9). This unexpected result may be attributed to the migration of biosurfactants to the 7


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oil–water interface from the aqueous phase because of the presence of excess electrolytes in

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the aqueous phase.29

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To transport B. subtilis to the target oil reservoir, we physically immobilized these

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bacteria in a hydrogel mesh of microgel particles. For production of microgel particles with

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uniformly immobilized B. subtilis, monodisperse emulsion drops were generated in a

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microcapillary-based microfluidic device (Fig. 5A). The aqueous mixture consisting of B.

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subtilis, PEGDA, NaSS, and photoinitiator was injected into the paraffin oil containing a

8

nonionic surfactant (Abil EM 90) by means of the coaxial jetting fluid phenomenon.

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Typically, the fluid thread that formed near the collection capillary tip pinched off into small

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drops because of hydrodynamic instability (Fig. 5B). The stability of dropping frequency led

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to formation of monodisperse emulsion drops. The coefficient of variation for our emulsion

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drops was less than 10%. After collection of the W/O emulsions, they were irradiated with

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UV light to polymerize them. Because B. subtilis is a few micrometers long, these bacteria

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can be immobilized within the hydrogel mesh whose mesh size is 5–10 nm in a swollen state

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in water19 (Fig. 6A–C). The efficiency of encapsulation of B. subtilis in the mesh of microgel

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particles was almost 100%, which is a key advantage of the microfluidic method. We found

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that the particle diameter of microgel particles in water increased by ~40% in comparison

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with the diameter of emulsion drops in paraffin oil (Fig. 6D). This finding can be attributed to

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the fact that the confined polymer network in the emulsion drop freely uncoiled in water, thus

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resulting in volume expansion.

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A core flooding experiment was performed on microgel particles with immobilized B.

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subtilis in a silica bead-filled model column. One pore volume (PV) of the microgel

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dispersion was injected into the glass beads after 2 PV of water was injected. Here, pore

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volume (PV) means total volume of void space in silica beads. 1 PV unit was calculated to be

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140 mL in our experiments. On the basis of the cumulative oil recovery with the PV of the

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injected fluid in the core flooding experiment, we could confirm that injection of the microgel

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particles significantly increased the oil recovery (Fig. 7A). Oil recovery was determined by

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measuring the amount of extracted oil from containing oil (original oil in place, OOIP; 119.8

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mL) in silica beads. Injection of water up to 2 PV led to 83.9-85.5 vol% (100.5-102.4 mL)

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release of OOIP from the silica glass beads. Additional 1 PV injection of the microgel fluid

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containing the B. subtilis-immobilized microgel particles increased the recovery of n-decane

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with the concentration-dependent manner. When the microgel particles were concentrated to 8


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5.0 wt%, the recovery of n-decane could be additionally enhanced over 7.1 vol % of OOIP

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(Fig. 7B). This means that 45.2 % (8.5 mL) of residual n-decane (18.8 mL) after water

3

injection was released.

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Additionally, to evaluate whether the viability of B. subtilis actually affect the oil

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recovery, we conducted the core flooding experiment using the microgel particles containing

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UV-killed B. subtilis cells. After UV-irradiation (254 nm) for over 1 h, the number of CFU of

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B. subtilis cells was decreased by 92%. Injection of 5 wt% microgel particles containing UV-

8

killed B. subtilis into silica bead led to the recovery of n-decane below 0.6 vol%. This

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recovery efficiency is quite comparable to the case of injecting the microgel particles

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containing living-B. subtilis. These results show that the use of microgel particles with

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immobilized B. subtilis successfully enhanced the oil recovery. Thus, the B. subtilis that was

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reliably protected by the hydrogel phase could be delivered to n-decane-filled porous media.

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Then, the biosurfactants produced by B. subtilis readily diffused out through the hydrogel

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mesh because their molecular size is several fold smaller than that of the hydrogel mesh pores

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in water. Under such reservoir conditions, the biosurfactants should preferably adhere to the

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n-decane captured in the silica glass beads to reduce the interfacial tension, thus eventually

17

facilitating recovery of the residual n-decane.

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There have been reports showing that a surfactant-producing strain of a Bacillus species

19

can enhance oil recovery.26,30,31 For studies on applicability of these bacteria to microbially

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enhanced oil recovery, the challenge is the issue of clogging during flooding.8,32 From the

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practical standpoint, the use of the microencapsulation technology not only enables protection

22

of microbes from the concentration reduction during flow in the medium but also allows

23

researchers to control the release rate of the incorporated microbes.10 For example, for active

24

delivery of capsules into an oil-bearing environment, a delivery protocol combining polymer

25

microcapsules with an organic-solvent stimulus has been developed.22 This approach

26

convincingly showed that the release of microbes from polystyrene microcapsules can be

27

triggered by solvation-induced capsule rupturing for oil recovery in porous media. Compared

28

to microcapsule-based microbe delivery, the method developed in the present study has an

29

advantage: we do not need to use such a trigger to induce the release of microbes from the

30

particles. As can be seen from the model diffusion experiments in Fig. 8, the small molecules

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that physically trapped in the mesh of microgel particles have a tendency toward slowly

32

diffusing out with no aids of external stresses. In fact, the diffusion kinetics are mainly 9


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determined by the mesh size. In the flooding experiment, therefore, we can expect that the

2

biosurfactants produced by microbes would simply diffuse out of the microgel particles and

3

show the ability to recover additional oil from oil-bearing structures.

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4. CONCLUSION

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Conventional methods of microbially enhanced oil recovery involve simple injection of

7

microbes or nutrients into the oil reservoir. In contrast, as proof of concept, we physically

8

immobilized microbesin microgel particles by the microfluidic method and tried to transport

9

these bacteria to the target reservoir. In this study, B. subtilis served as a model microbe that

10

can produce biosurfactants. We confirmed empirically that B. subtilis reduces the interfacial

11

tension between oil and water. In the flooding experiment, we demonstrated that injection of

12

microgel particles (on which B. subtilis is immobilized) after the water flooding may enhance

13

oil recovery over 7 vol%, which corresponds to more than 45 vol% recovery of the residual

14

oil from a model reservoir column packed with silica beads. This finding is comparable to the

15

result of flooding water only. Such improved oil recovery is possible because B. subtilis

16

produces biosurfactants that readily diffuse out of the microgel particles through the hydrogel

17

mesh in the reservoir, thus facilitating emulsification of oil in water during oil recovery.

18

Consequently, the proposed technology for fabrication of microgel particles with

19

immobilized microbes is expected to be applied to microbially enhanced oil recovery in

20

permeable porous media.

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AUTHOR INFORMATION

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Corresponding Author: Prof. Jin Woong Kim

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*E-mail: [email protected]

25

*Telephone: +82-31-400-5499

26

*Address: Department of Bionano Technology and Department of Applied Chemistry,

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Hanyang University, Ansan 15588, Republic of Korea.

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Corresponding Author: Dr. Hyun-Tae Kim

29

*E-mail: [email protected]

30

*Telephone: +82-42-868-3216.

31

*Address: Korea Institute of Geoscience and Mineral Resources, Daejeon 34132, Republic of 10


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Korea.

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The authors declare no competing financial interest.

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ACKNOWLEDGMENTS

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The work was carried out with financial support from the Korea Institute of Geosciences and

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Mineral Resources and by the National Research Foundation of Korea (NRF) grant funded by

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the Korea government (MSIP) (No. 2008-0061891 and No. 2016R1A2B2016148). H. A. Son

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and S. K. Choi equally contributed to this work.

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Graphical Abstract

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Figure 1. A schematic diagram of the experimental apparatus for microbially enhanced oil

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recovery.

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Figure 2. Changes in the interfacial tension of n-decane-water with changing the

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concentration of B. subtilis and E. coli in the aqueous phase. The initial concentration of B.

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subtilis and E. coli in the aqueous phase was tuned to be 0.9 g/L. The deviation in the values

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of interfacial tensions came from pendant drop profiles.

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Figure 3. Changes in the interfacial tension of n-decane–water interfaces as a function of (A)

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pH and (B) NaCl concentration in the presence of B. subtilis in the aqueous phase. The

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concentration of B. subtilis in the aqueous phase was set to 0.9 g/L in (A) and (B), and pH

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was adjusted to 8 in (B).

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Figure 4. Interfacial activation of Bacillus subtilis in n-decane emulsions. (A) Immediately

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after emulsification and (B) after 7-day storage at room temperature. Sample 1 corresponds to

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the n-decane emulsion without B. subtilis (pure water). Samples 2–5 correspond to n-decane

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emulsions containing B. subtilis: 0, 2, 5, or 10 wt% NaCl, respectively, at pH 4. Samples 6–9

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are the n-decane emulsions containing B. subtilis: 0, 2, 5, and 10 wt% NaCl, respectively, at

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pH 8.

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Figure 5. (A) A schematic of a coaxial jetting microfluidic capillary device. (B) A bright-

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field micrograph showing formation of monodisperse microgel precursor emulsion drops

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containing B. subtilis.

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Figure 6. (A) A bright-field micrograph of B. subtilis in the aqueous phase. Bright-field

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micrographs of (B) microgel precursor emulsion drops and (C) microgel particles with

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immobilized B. subtilis. (D) Particle size distributions before and after microgel formation.

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Figure 7. (A) Overall oil recovery as a function of pore volume for injection of water and

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microgel particles with immobilized B. subtilis. (B) Cumulative oil recovery immediately

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after injection of microgel particles with B. subtilis. The injection fluid contained 0.05 wt%

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(), 1.5 wt% (), 5.0 wt% (◆) microgel particles with living B. subtilis and 5.0 wt% (▲)

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microgel particles with UV-killed B. subtilis.

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Figure 8. (A) Releasing kinetics of fluorescein sodium salt (FSS, 376.27 g/mol) from

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microgel particles at 25°C. FSS was physically encapsulated in PEGDA microgel particles:

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700 g/mol PEGDA (~75 Å calculated mesh size, --), 6000 g/mol PEGDA (~290 Å

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calculated mesh size, --). Fluorescence microscopy images of FSS-loaded PEGDA

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microgel particles: 700 g/mol PEGDA after storage for 1 day (B) and 11 days (C), 6000

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g/mol PEGDA after storage for 1 day (D) and 11 days (E) in water.

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Immobilized Microgel Particles for Enhanced Oil Recovery - American

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