RESEARCH PAPER
A plug-based microfluidic system for dispensing lipidic cubic
phase (LCP) material validated by crystallizing membrane
proteins in lipidic mesophases
Liang Li • Qiang Fu • Christopher A. Kors •
Lance Stewart • Peter Nollert • Philip D. Laible •
Rustem F. Ismagilov
Received: 5 August 2009 / Accepted: 22 September 2009

Springer-Verlag 2009
Abstract This article presents a plug-based microfluidic
system to dispense nanoliter-volume plugs of lipidic cubic
phase (LCP) material and subsequently merge the LCP
plugs with aqueous plugs. This system was validated by
crystallizing membrane proteins in lipidic mesophases,
including LCP. This system allows for accurate dispensing
of LCP material in nanoliter volumes, prevents inadvertent
phase transitions that may occur due to dehydration by
enclosing LCP in plugs, and is compatible with the tradi-
tional method of forming LCP material using a membrane
protein sample, as shown by the successful crystallization
of bacteriorhodopsin from Halobacterium salinarum.
Conditions for the formation of LCP plugs were charac-
terized and presented in a phase diagram. This system was
also implemented using two different methods of intro-
ducing the membrane protein: (1) the traditional method of
generating the LCP material using a membrane protein
sample and (2) post LCP-formation incorporation (PLI),
which involves making LCP material without protein,
adding the membrane protein sample externally to the LCP
material, and allowing the protein to diffuse into the LCP
material or into other lipidic mesophases that may result
from phase transitions. Crystals of bacterial photosynthetic
reaction centers from Rhodobacter sphaeroides and
Blastochloris viridis were obtained using PLI. The plug-
based, LCP-assisted microfluidic system, combined with
the PLI method for introducing membrane protein into
LCP, should be useful for minimizing consumption of
samples and broadening the screening of parameter space
in membrane protein crystallization.
Keywords Droplet
Plugs
Lipidic cubic phase

Membrane protein
Protein crystallization
1 Introduction
This article describes a novel procedure to manipulate
viscous lipidic cubic phase (LCP) material in a plug-based
microfluidic system for crystallizing membrane proteins.
Obtaining high quality crystals of membrane proteins is
important for determining their structures (Ostermeier and
Michel 1997). Purification and crystallization of membrane
proteins in detergent solutions may lead to reduced activity
and poor stability of the proteins. On the other hand, using
a lipidic mesophase, such as LCP, is an important route to
the crystallization of membrane proteins because the lipidic
mesophase provides an environment similar to the natural
environment of membrane proteins (Caffrey 2000). Crys-
tallization in LCP material has been shown to be an
important approach for obtaining high quality crystals of
Electronic supplementary material The online version of this
article (doi:10.1007/s10404-009-0512-8) contains supplementary
material, which is available to authorized users.
L. Li
Q. Fu
R. F. Ismagilov (&)
Department of Chemistry and Institute for Biophysical
Dynamics, The University of Chicago, 929 East 57th Street,
Chicago, IL 60637, USA
e-mail: r-ismagilov@uchicago.edu
C. A. Kors
P. D. Laible
Biosciences Division, Argonne National Laboratory, 9700 South
Cass Ave., Argonne, IL 60439, USA
L. Stewart
deCODE biostructures, Accelerated Technologies Center
for Gene to 3D Structure, 7869 NE Day Rd. W,
Bainbridge Island, WA 98110, USA
P. Nollert
Emerald BioSystems, Inc., 7869 NE Day Rd. W,
Bainbridge Island, WA 98110, USA
123
Microfluid Nanofluid
DOI 10.1007/s10404-009-0512-8

membrane proteins, as demonstrated by the recently
determined structures for two G protein-coupled receptors
(Cherezov et al. 2007; Jaakola et al. 2008).
Current developments in LCP-based microscale protein
crystallization include the development of robotic systems
that allow for accurate handling of small amounts of LCP
material (Cherezov and Caffrey 2006; Peddi et al. 2007;
Perry et al. 2009) and the development of a sparse matrix
screening kit, which involved pre-mixing different pre-
cipitants with monoolein to make lipidic sponge phase
material (Wohri et al. 2008). Yet, crystallization in LCP
material has still not been widely applied to crystalliza-
tion. One reason is because LCP material has high vis-
cosity, which makes it difficult to dispense with high
accuracy using traditional tools for dispensing liquids.
Furthermore, because dehydration of LCP material using
conventional dispensing methods (Cherezov et al. 2004;
Nollert 2002) can lead to phase transitions, current
methods for working with small volumes of LCP material
must be performed at high humidity to account for water
loss. As each experimental setup may require a specific
humidity, these methods can be complicated, take a long
time, and, if carried out in parallel for a variety of con-
ditions, alter the desired composition of the crystallization
cocktail.
Recently, a microfluidic system using pneumatic valves
which formed LCP material on-chip at volumes below
20 nl was developed for crystallization of membrane
proteins (Perry et al. 2009). However, this system used
PDMS to create the pneumatic valves, and PDMS may
cause unwanted evaporation and loss of chemicals in
crystallization trials, such as lipids (Toepke and Beebe
2006). Like the pneumatic valve system, the plug-based
microfluidic system (Song et al. 2003, 2006a) that has
been developed for protein crystallization (Lau et al. 2007;
Shim et al. 2007; Zheng et al. 2005; Zheng et al. 2003)
accurately handles nanoliter volumes of viscous fluids in
microfluidic channels. The plug-based system has been
demonstrated with clotted blood (Song et al. 2006b) and
solutions containing nanoparticles (Shestopalov et al.
2004). Unlike the pneumatic valve system, using plug-
based microfluidics eliminates loss of chemicals in crys-
tallization trials as well as unwanted evaporation that can
lead to phase transitions. Moreover, the plug-based
microfluidic system offers the ability to simultaneously
handle small volumes of different precipitants and
manipulate their concentrations with high precision by
using the hybrid method (Li et al. 2006).
In this article, we extend the plug-based microfluidic
system to manipulation of LCP material. We demon-
strated the formation of nanoliter-volume LCP plugs and
the ability to merge those plugs into aqueous plugs. We
then validated the developed system by crystallizing
membrane proteins. We introduced membrane protein
samples into LCP by using two different methods: (1) the
traditional method of generating the LCP material using a
membrane protein sample (Fig. 1a) and (2) post LCP-
formation incorporation (PLI), in which a membrane
protein sample is combined with the LCP material and
allowed to diffuse into the LCP material (Fig. 1b) or into
other lipidic mesophases, which may result from phase
transitions induced by certain precipitants, such as Jeff-
amine M-600 (Wadsten et al. 2006; Wohri et al. 2008).
We successfully crystallized bacteriorhodopsin (BR) from
Halobacterium salinarum using the traditional method,
and we successfully crystallized the bacterial photosyn-
thetic reaction center (RC) from two strains of Rhodob-
acter sphaeroides-a carotenoid-containing strain (Davis
et al. 1988) and a carotenoidless strain (Theiler et al.
1984)-as well as RC from Blastochloris viridis, using
PLI.
2 Materials and methods
2.1 Chemicals
All solvents and salts purchased from commercial sources
were used as received unless otherwise stated. Lauryl-
dimethylamine n-oxide (LDAO) and n-octyl-b-glucopy-
ranoside (OG) were purchased from Anatrace (Maumee,
OH). 9-Monoolein was obtained from Nu-Check Prep, Inc.
(Elysian, MN). (tridecafluoro-1,1,2,2,-tetrahydrooctyl)-1
trichlorosilane was obtained from United Chemical
Fig. 1 Schematic showing two different methods to crystallize
membrane proteins in LCP material. a Traditional method: the LCP
material is made with protein. b Post LCP-formation incorporation
(PLI): the LCP material is made without protein, then a membrane
protein sample is added externally and allowed to diffuse into the
LCP material for crystallization. * In experiments with certain
precipitants, the LCP material may transform into other lipidic
mesophase materials, as a result of a phase transition induced by the
precipitants
Microfluid Nanofluid
123

Technologies, Inc. (Bristol, PA). Poly(dimethylsiloxane)
(Sylgard 184 Silicone Elastomer kit) was obtained from
Dow Corning (Midland, MI). FC-40, a mixture of per-
fluoro-tri-n-butylamine and perfluoro-di-n-butylmethyl-
amine, and FC-70, perfluorotripentylamine, were obtained
from 3 M (St. Paul, MN).
2.2 Equipment
Spectra were analyzed using a UV–visible spectrometer
purchased from Agilent (Santa Clara, CA). Teflon tubing
(O.D. 250 lm, I.D. 200 lm) and Teflon tubing (O.D.
250 lm, I.D. 100 lm) were purchased from Zeus
(Orangeburg, SC). Teflon tubing (O.D. 750 lm, I.D. 300 lm)
was obtained from Weico Wire & Cable (Edgewood, NY).
Standard wall glass tubing was obtained from Chemglass
(Vineland, NJ). Gastight syringes were obtained from
Hamilton Company (Reno, NV).
2.3 Fabricating PDMS devices
All microfluidic devices were fabricated from poly(dimeth-
ylsiloxane) (PDMS). Microchannels with rectangular cross
sections were fabricated by using rapid prototyping
(Duffy et al. 1998). The channel walls were modified
with (tridecafluoro-1,1,2,2-tetrahydrooctyl)-1-trichlorosi-
lane to increase their hydrophobicity and fluorophilicity
(Roach et al. 2005).
2.4 Assembling the microfluidic system
The microfludic system consisted of two devices: one
was a flow-focusing device that generated LCP plugs
(Fig. 2a, b), and the other was a merging device in which
aqueous plugs were formed and then merged with LCP
plugs (Fig. 2c). A piece of Teflon tubing (O.D. 250 lm,
I.D. 100 lm, 7–10 cm in length) was used to connect the
two devices. To prevent the LCP plugs from contacting
the PDMS channels, one end of the tubing was inserted
through the outlet flush with the cross junction of the
flow-focusing device (Fig. 2b), and the other end was
inserted flush with the junction where aqueous plugs were
formed via the fluorocarbon inlet of the merging device
(Fig. 2c). At the junction, the Teflon tubing is wetted
preferentially by FC-70, allowing LCP plugs to be
formed and transported. Through the outlet of the
merging device, another piece of Teflon tubing (O.D.
550 lm, I.D. 400 lm, *20 cm in length) was inserted
flush with the junction where aqueous plugs were formed
(Fig. 2c). All the gaps between the PDMS channels and
the Teflon tubing were sealed with wax to prevent
leakage.
2.5 Forming LCP plugs in the flow-focusing device
LCP material was first made by mixing monoolein and
Millipore water (w:w, 3:2) using an LCP mixer obtained
from Emerald BioSystems, Inc. (Nollert 2002). Once the
LCP material was formed, it was transferred into a syringe,
and the mixer was disassembled. The syringe containing
the LCP material was coupled to a 27-gauged syringe
needle, which was connected to a piece of Teflon tubing
(I.D. 300 lm, O.D. 750 lm, *5 cm in length). The LCP
material was then transferred into the Teflon tubing. Once
the tubing was filled, it was detached from the needle and
was then attached to another 10 ll Hamilton glass syringe
prefilled with FC-40. The tubing was then attached to the
PDMS device, ready for forming LCP plugs.
2.6 Conditions for formation of LCP plugs
To determine the conditions for the formation of LCP
plugs, the flow-focusing device was used without being
connected to the merging device. A piece of Teflon tubing
(O.D. 250 lm, I.D. 100 lm) was inserted flush with the
cross junction via the outlet and the gap between the
PDMS channel, and then the Teflon tubing was sealed
with wax. At flow rates of the carrier fluid (FC-70)
between 2 and 3 ll/min, LCP plugs formed successfully
(Fig. 2e). All the solutions and LCP material in these
experiments were loaded in glass syringes, and the flow
rates were controlled by using syringe pumps from Har-
vard Apparatus.
2.7 Consistency of plug formation
To determine the consistency of the volume of the LCP
plugs, the length of the plugs was monitored. In the same
experiment, we also monitored the distance between plugs
to determine the stability of flow in the device. In this
experiment, the flow rates of LCP material and FC-70 were
kept at 0.2 and 2 ll/min, respectively, and images of plugs
were taken at a fixed position at nine different time points:
0, 2, 5, 9, 13, 16, 18, 23, and 28 min. For each time point,
images of five different plugs were taken. The length of
plugs in all images was then measured, as was the distance
between two adjacent plugs (Fig. 3).
2.8 Stability of LCP plugs
To determine whether LCP material maintained its phase
during formation of plugs, LCP plugs were formed,
transported in Teflon tubing (100 lm I.D.), and imaged
under cross-polarization. The flow rates were 0.3 ll/min
for the LCP material and 2 ll/min for FC-70. After LCP
plugs formed, the Teflon tubing was disconnected from the
Microfluid Nanofluid
123

device and then the tubing was sealed at both ends. Images
of plugs in the tubing were taken within 1 h under cross-
polarized light (Fig. 4). We took images at two different
angles of cross-polarized light to eliminate the possibility
that background birefringence from the Teflon tubing could
have hid any birefringence caused by the LCP material.
2.9 Merging LCP plugs with aqueous plugs
To determine whether LCP plugs could successfully merge
with aqueous plugs in the microfludic system, streams of
50% Polyethyleneglycol(PEG)-8000 and Millipore water
were used as the precipitant stream and the protein stream,
respectively, and LCP plugs were formed by using LCP
materials made from monoolein and water, as described
above. The flow rate of FC-70 was maintained at 3 ll/min
throughout the experiment. A phase diagram was generated
by using the total flow rate of aqueous solutions between 1
and 4 ll/min and using the flow rate of the LCP material
between 0.1 and 0.3 ll/min (Fig. 2f–i). At a given total
flow rate, each of the aqueous solutions flowed at a rate
equal to half of the total flow rate.
Fig. 2 Plug-based microfluidic system for crystallization of mem-
brane proteins within lipidic mesophases. a A schematic of the
microfluidic system. Small LCP plugs (*1 nl) were formed in a
PDMS flow-focusing device using fluorinated carbon (FC) as a carrier
fluid. The LCP plugs were transported in Teflon tubing, and then they
merged downstream with the streams of protein and precipitant
cartridges in a PDMS device to form LCP-containing aqueous plugs
(*80 nl). The stream of protein was added only when LCP material
was made using PLI (Fig. 1b). Upon merging with plugs containing
certain precipitants, the LCP material may undergo phase transition to
form another lipidic mesophase material, commonly known as a
sponge phase (Wadsten et al. 2006). The plugs of the crystallization
trials were stored and incubated at 23C in Teflon tubing to allow
crystals to grow. b A micrograph showing that LCP plugs formed in
the flow-focusing device. c A micrograph showing that LCP plugs
successfully merged with precipitant and protein solutions. LCP plugs
are delineated by dashed white lines. d A micrograph showing that the
plugs to the right of the air bubble did not contain red solution,
indicating the absence of cross contamination of aqueous plugs
separated by air bubbles (Li et al. 2006). e A phase diagram showing
the working range of the flow rates of FC and LCP material that are
required for reliably forming LCP plugs. The solid squares indicate
reliable formation of LCP plugs, and the open squares indicate failure
to form LCP plugs. f A phase diagram showing the flow rates of the
LCP material and the aqueous flow rates that are required for reliable
merging of LCP plugs with aqueous plugs. g A micrograph showing
redundant merging caused by low aqueous flow rate and high LCP
flow rate, indicated by solid triangles in the phase diagram. h A
micrograph showing reliable merging at the working flow rates,
indicated by the solid squares in the phase diagram. i A micrograph
showing insufficient merging at high aqueous flow rate and low LCP
flow rate, indicated by the open circle in the phase diagram
Microfluid Nanofluid
123

2.10 Forming cartridges for the precipitant stream
To determine whether cartridges can be implemented in
the microfludic system, Millipore water was used as the
protein stream, and a cartridge, consisting of 200 nl pre-
cipitant-mimicking plugs alternated with 50 nl air bub-
bles, was used as the precipitant stream. The cartridge
contained 24 precipitant-mimicking plugs, of which 12
were colorless plugs, comprising conditions 1–12 from
Hampton Research Index Screening Kit, and 12 were red
plugs, containing 0.1 M Fe(SCN)3 solution. The cartridge
was prepared by alternating colorless plugs and red plugs.
In the experiment, the flow rates of FC-70, the LCP
material, the protein stream, and the precipitant stream
were maintained at 3, 0.3, 0.5, and 1.5 ll/min, respec-
tively. Images were recorded during and after the merging
process (Fig. 2c, d).
2.11 Preparing samples of membrane proteins
Halobacterium salinarum S9 was grown using the pub-
lished protocol (Cline and Doolittle 1987), and samples of
bacteriorhodopsin (BR) were obtained by purification from
membranes using the reported procedures (Nollert 2004)
with separation proceeding on a GE Healthcare HiLoad
TM
26/60 Superdex
TM
75 Prep Grade column.
Carotenoid-containing and carotenoidless strains of
R. sphaeroides were grown semi-aerobically in the dark in
YCC Medium (Taguchi et al. 1992), and samples of poly-
histidine-tagged RCs from R. sphaeroides were obtained by
purification with minimal light exposure to a purity marked
with optical absorbance ratio of A800/A280 * 1.5 as previ-
ously described (Pokkuluri et al. 2002). Anion exchange
chromatography (Tiede et al. 1996) was used to further
improve purity of the samples and to increase crystalliza-
tion reproducibility, bringing optical absorbance ratios of
A800/A280 to between 1.2 and 1.3.
Polyhistidine-tagged carotenoidless RCs were pro-
duced using a new expression strategy. A recombinant
strain was created by mobilizing the expression vector
pRKHTMHBgl (Pokkuluri et al. 2002), which carried a
wild-type version of the gene encoding the L subunit and a
modified version of the gene encoding the M subunit, into
the host R. sphaeroides strain R26.1 (Theiler et al. 1984)
via conjugation using the donor E. coli strain S17-1 (Simon
et al. 1983). The new strain, R26.1[pRKHTMHBgl],
simultaneously expressed tagged and untagged versions of
the RC from R. sphaeroides. These two types of RCs were
easily separated by metal affinity chromatography after
being solubilized by using LDAO from intracytoplasmic
membranes of the host strain.
2.12 Crystallizing BR
LCP material was made using the method described above
(Nollert 2002), with the exception that Millipore water was
replaced by a solution of BR sample. 30 mg monoolein and
20 ll BR, at a concentration of 22.5 mg/ml, were con-
sumed to make the LCP material. Crystallization trials
were set up in the microfludic system, which consisted of a
flow-focusing device and a merging device bearing one
aqueous inlet (Fig. 5c I). In the flow-focusing device, the
flow rate of the carrier fluid, FC-70, was 3 ll/min, and the
flow rate of the LCP material was 0.2 ll/min. Although
the presence of detergents in a protein sample have an
effect on LCP material (Ai and Caffrey 2000), we were still
able to form LCP plugs in the presence of detergent
(LDAO). Precipitant cartridges were used as the aqueous
stream in the merging device. The cartridges consisted of
200 nl precipitant plugs alternated with 50 nl air bubbles.
Fig. 3 LCP plugs formed consistently in the flow-focusing device.
The flow rate of LCP material was 0.2 ll/min and the flow rate of FC-
70 was 2 ll/min. The plug length (solid squares), which was directly
correlated to the volume, was *150 lm over more than 25 min. The
standard deviation of the plug length was 8%, indicating that the
standard deviation for the plug volume was 8%. The space between
plugs (solid triangles) was *800 lm over the same time period, with
16% variation
Fig. 4 LCP material maintained its phase during plug formation.
Micrographs indicated the absence of birefringence of the LCP
material at two different angles of cross-polarized light. Only
background pattern, raised from the Teflon tubing, was observed
Microfluid Nanofluid
123

Two cartridges, each containing 25 conditions from Crystal
Screen kit (Hampton Research), were prepared as previ-
ously reported (Li et al. 2006), resulting in 50 total con-
ditions. The cartridges were flowed at a rate of 2.0 ll/min.
Once the crystallization trials were set up, the Teflon
tubing containing the trials was detached from the network
and sealed in a piece of glass tubing prefilled with
FC-70. The trials were then incubated at 23C in the
dark.
2.13 Crystallizing carotenoid-containing RC
LCP material was made using the method described
above (Nollert 2002). 30 mg monoolein and 20 ll Milli-
pore water were used to make the LCP material.
Crystallization trials were set up in the microfluidic sys-
tem, which included a flow-focusing device and a merg-
ing device bearing three aqueous inlets (Fig. 5d I): the
first for precipitant cartridges, the second for the buffer
(0.05% (w/v) LDAO, 10 mM Tris pH 7.8), and the third
for the protein sample (20 mg/ml in 0.05% (w/v) LDAO,
10 mM Tris pH 7.8). The precipitant cartridges were
made in the same way as described above, except that a
different kit, provided by Emerald Biosystems, was used
to prepare 48 different conditions (Table S1). The carrier
fluid, FC-70, was flowed at a rate of 3 ll/min; the flow
rate of LCP material was 0.3 ll/min. The flow rates of the
precipitant cartridge, the buffer, and the protein sample
were 1.5, 0.1, and 0.5 ll/min, respectively. The trials
were incubated as described above. Crystals began to
appear within a week.
Fig. 5 a A schematic showing the traditional method for introducing
protein into LCP material. b A schematic showing PLI, a method of
forming LCP material without protein, adding the membrane protein
sample externally to the LCP material, and allowing the protein to
diffuse into the LCP material. c Crystallization of BR using the
traditional method. c, I—A schematic showing the experimental setup
for crystallization. c, II—A plug with crystals (in dark spots) of BR
(from H. salinarum) obtained when BR was premixed in LCP
material. d, e, f Crystallization of three proteins using PLI. (I)
Schematics showing the experimental setups for crystallization using
PLI. For the carotenoid-containing RC from R. sphaeroides (d, I),
additional buffer was added to the precipitant cartridge and protein
streams. For the carotenoidless RC from R. sphaeroides (e, I) and RC
from B. viridis (f, I), no additional buffer was added to the precipitant
cartridge and protein streams. (II) Plugs with crystals obtained using
PLI of (c, II) carotenoid-containing RC, (d, II) carotenoidless RC, and
(e, II) RC (B. viridis). (III) Diffraction-quality crystals were obtained
from proteins grown in lipidic mesophase material. c, III—A crystal
of carotenoid-containing RC diffracted X-ray to *3.5 A˚ (ring
indicates 3.6 A˚ resolution). d, III—A crystal of carotenoidless RC
grown in lipidic mesophase material diffracted X-ray to *2.5 A˚ (ring
indicates 3.0 A˚ resolution). e, III—A crystal of RC (B. viridis)
diffracted X-ray to *2.8 A˚ (ring indicates 3.0 A˚ resolution)
Microfluid Nanofluid
123

2.14 Crystallizing carotenoidless RC
The same procedure that was followed for crystallizing
carotenoid-containing RC was followed for crystallizing
carotenoidless RC. In this case, however, the merging
device contained only two aqueous inlets (Fig. 5e I): one
for precipitant cartridges, flowing at a rate of 1.8 ll/min,
and the other for the protein sample (20 mg/ml in 0.05%
(w/v) LDAO, 10 mM Tris pH 7.8), flowing at a rate of
0.2 ll/min. The same screening kit, provided by Emerald
Biosystems, as was used for crystallizing carotenoid-con-
taining RC was also used here. The trials were incubated as
described above. Crystals began to appear within 2 days.
2.15 Crystallizing RC from B. viridis
The same procedure that was followed for crystallizing
carotenoidless RC was followed for crystallizing RC from
B. viridis (Fig. 5f I). The protein sample was 26 mg/ml in
0.08% (w/v) LDAO, 50 mM Na2PO4–NaH2PO4, pH 6.0.
The same screening kit and incubation procedures were
used. Crystals began to appear within a week.
2.16 Preparing crystals for X-ray diffraction
Crystallization trials were checked by using a stereoscope
under minimal light. The same procedure as previously
reported (Li et al. 2006) was followed, and plugs con-
taining crystals were flowed into a droplet of the mother
liquor sitting in a microwell. No cryoprotectant was nee-
ded. Crystals, together with the lipidic mesophase material,
were looped from droplets directly and were then flash
frozen in liquid nitrogen.
2.17 X-ray diffraction and data processing
X-ray diffraction was performed at GM/CA Cat station 23
ID-B of the Advanced Photon Source (Argonne National
Laboratory). The data were processed in HKL2000
(Otwinowski and Minor 1997).
3 Results and discussion
3.1 LCP material reliably formed nanoliter-volume
plugs
First, we demonstrated the compatibility of LCP material
with the plug-based microfluidic system (Fig. 2). LCP
plugs were first formed in a microfluidic stream (Fig. 2a).
The working flow rates in these experiments are shown in a
phase diagram (Fig. 2e). The minimum working flow rate
was limited by the viscosity of the LCP material. At flow
rates of FC of 2 ll/min or higher, the cross flow of the FC
provided shear high enough to cut the viscous LCP stream
into nanoliter-volume plugs (Fig. 2b). The maximum
working flow rate was limited by the maximum pressure
the device could withstand, *0.4 MPa (McDonald et al.
2000). At flow rates of FC of 4 ll/min or higher, the
pressure on the device was greater than 0.4 MPa. The
volume of the LCP plugs, *1 nl, was consistent over time
with a standard deviation of 8% (Fig. 3). We checked the
stability of LCP material in the carrier fluid in a separate
experiment, and the LCP material maintained its phase
while forming plugs (Fig. 4).
3.2 LCP plugs successfully merged with aqueous plugs
to create crystallization trials without cross
contamination
When the LCP plugs came in contact with the aqueous
plugs, they merged with the aqueous plugs, presumably to
minimize interfacial energy due to the hydrophilic surface
of the LCP material. In the presence of certain precipitants,
the LCP material in the merged plugs may undergo a phase
transition to form another lipidic mesophase, commonly
known as a sponge phase (Fig. 2a, d). The efficiency of
merging of LCP plugs with aqueous plugs for the crystal-
lization trials was governed by the flow rate of the LCP
material upstream and the aqueous flow rate at which the
plugs for crystallization trials were formed (Fig. 2f). To
describe this effect, we defined the frequency f1 as the
number of LCP plugs that were formed upstream per sec-
ond and the frequency f2 as the number of aqueous plugs of
crystallization trials that were formed per second. The
ratio, N = f1/f2, denotes the number of LCP plugs in each
aqueous plug and defines the efficiency of merging. The
lower limit of N is 1, because every aqueous plug must
merge with at least one LCP plug, and the upper limit of N
is set by the requirement that a single aqueous plug must
accommodate all the LCP plugs formed upstream. When
the flow rate of LCP material and the aqueous solution was
moderate (flow rate of LCP material = 0.1 or 0.2 ll/min;
aqueous flow rate = 1, 2, or 3 ll/min), N was greater than
1 but less than 8, and reliable merging was observed
(Fig. 2f, solid squares, h). When the flow rate of the LCP
material was too high (0.3 ll/min or higher), f1 increased
and so did N, up to values over 8. This resulted in redun-
dant merging, in which more than one LCP plug merged
with each aqueous plug (Fig. 2f, solid triangles, and g).
When the aqueous flow rate was too high (over 4 ll/min),
f2 increased, and N dropped below 1. At low LCP flow
rates, the LCP material could not reliably merge with every
aqueous plug, resulting in insufficient merging (Fig. 2f,
open circles, and i), and at higher LCP flow rates redundant
merging was again observed.
Microfluid Nanofluid
123

Under the working conditions described above, the LCP
plugs preferentially merged with the aqueous plugs, and
each merged plug constituted a crystallization experiment
with no cross contamination between conditions (Fig. 2d).
This method allowed for sparse matrix screening of crys-
tallization conditions with LCP material.
3.3 Model membrane proteins crystallized using the
plug-based LCP-assisted microfluidic system
Having demonstrated the compatibility of the plug-based
microfluidic system with LCP material, we then validated
the system by crystallizing proteins (Table 1) using the two
different methods of introducing membrane protein into
LCP material (Fig. 1).
To test the compatibility of our system with method 1
(Fig. 5a), we selected BR from H. salinarum, and we
prepared the LCP material by directly mixing monoolein
with a solution of BR (3:2 (w/w) ratio). In this experi-
ment, the aqueous stream was a single stream, containing
an array of precipitant plugs (Fig. 5c I). Conditions were
screened using a commercial kit (Crystal Screen from
Hampton Research) and crystals were obtained (Fig. 5d) in
16 different conditions (Table 1, the specific conditions are
listed in Table S2). Thus, the traditional method for making
LCP material with protein was successfully implemented
in the plug-based system. Although BR crystals were too
small to be characterized by X-ray diffraction, the crystals
were identified by the color of the protein and the shape of
the potential crystal (Fig. 5c II).
To demonstrate PLI, three target proteins were screened
against conditions formed with various combinations of
Jeffamine M-600 and (NH4)2SO4 (Table S1), 48 different
conditions in total. The targets were membrane proteins
with known crystallization conditions that included RCs
from two bacterial species: R. sphaeroides and B. viridis.
In the experimental configurations for all targets, proteins
could be conveniently injected as an aqueous stream lam-
inar with the stream containing the precipitant cartridge
(Fig. 5d I, e I, f I). The precipitant cartridge contained an
array of aqueous precipitant plugs (*200 nl in volume)
separated by air bubbles (*50 nl in volume); each pre-
cipitant plug contained unique conditions for protein
crystallization. In these crystallization experiments, each
precipitant plug formed three to four smaller aqueous plugs
that then merged with the LCP plugs; the plugs formed in
this way from a single precipitant plug constituted replicate
crystallization trials of a given condition. Thus, with a
protein sample of *3 ll, 48 different conditions, with 3–4
replicates each, could be set up within 6 min.
Some of the LCP plugs, upon merging with aqueous plugs
containing precipitant and protein, underwent a phase tran-
sition to another lipidic mesophase (Cherezov et al. 2006;
Cherezov et al. 2001). Crystals grew directly in the new
lipidic mesophase material (Fig. 5d II, e II, f II). Crystals of
carotenoid-containing RC from R. sphaeroides 2.4.1
formed (Fig. 5d II) in the mesophase within a week in 3 of 48
conditions (Table 1, the specific conditions are listed in
Table S1) and diffracted X-rays to *3.5 A˚ (Fig. 5d III).
Crystals of caroteniodless RC from R. sphaeroides R26
formed (Fig. 5e II) in the lipidic mesophase material within
2 days in 16 of 48 conditions (Table 1, the specific condi-
tions are listed in Table S1) and diffracted X-rays to*2.5 A˚
(Fig. 5e III). Crystals of RC from B. viridis formed (Fig. 5f II)
in the lipidic mesophase material within a week in 6 of 48
conditions (Table 1, the specific conditions are listed in
Table S1) and diffracted X-rays to *2.8 A˚ (Fig. 5f III).
Crystals of carotenoidless RC from R. sphaeroides R26
obtained by using PLI (Fig. 5e III) belonged to P42212 with
the unit cell parameter of a = b = 101.0 A˚, c = 238.4 A˚;
a = b = c = 90. These results were similar to those
obtained previously by forming LCP material with protein
(Katona et al. 2003), validating the PLI method. The
consistency in the unit cell parameter between the results
obtained here and those reported previously may imply that
crystallization occurred after RC diffused into the lipidic
mesophase when PLI was used. That is, the same crystal
morphology may be due to crystallization in both cases
sharing the same mechanism.
4 Conclusion
We developed a plug-based microfluidic system that
accurately dispenses nanoliter volumes of lipidic cubic
phase (LCP) material. LCP material formed plugs of
nanoliter volumes with 8% deviation in length among
plugs. Conditions for the formation of LCP plugs were
Table 1 Summary of crystallization experiments using membrane protein targets in the plug-based LCP-assisted microfluidic system
Protein Source M.W. (kDa) Detergent
(mM)
Protein
(mg/ml)
Crystallization hits
out of 48 precipitants
Bacteriorhodopsin H. salinarum 27 40 12–30 16
Carotenoid-containing reaction center R. sphaeroides 2.4.1 derivative 100 2 10–25 3
Carotenoidless reaction center R. sphaeroides R26 100 2 *20 16
Reaction center with bound cytochrome B. viridis 135 3.5 *20 6
Microfluid Nanofluid
123

characterized and presented in a phase diagram. We vali-
dated this system by successfully screening conditions for
crystallization of membrane proteins; each crystallization
plug constituted an individual experiment under different
conditions. The system we have described is compatible
with the traditional method of pre-forming LCP material
using a membrane protein sample, as well as with PLI, a
method of forming LCP material without protein, adding
the membrane protein sample externally to the LCP
material, and allowing the protein to diffuse into the LCP
material. Because PLI relies on diffusion of the protein into
the LCP material, it may be limited by any factors that
limit this diffusion, such as the size of the protein. We will
test a batch of membrane proteins with different sizes of
hydrophilic areas in future studies. Nevertheless, this
alternative approach eliminated the step of preparing LCP
material individually for every protein target studied. This
alternative method minimizes sample consumption because
the protein samples can be filled into syringes without loss
in dead volumes. The volumes used in this system could
potentially be scaled down further: Using the current
device with the pressure limit of 0.4 MPa, LCP plugs as
small as 200 pL should be formed at a total flow rate of
0.4 ll/min in a 7 cm long piece of Teflon tubing with
60 lm I.D. Furthermore, by adding extra inlets for aqueous
streams, the system could be made compatible with more
complicated methods of screening, such as additive meth-
ods and those using ligands. This method should comple-
ment current developments in LCP-based microscale
protein crystallization (Cherezov and Caffrey 2006; Peddi
et al. 2007; Perry et al. 2009).
Acknowledgments This work was supported through Accelerated
Technologies Center for Gene to 3D Structure (ATCG3D) funded by
the National Institute of General Medical Sciences (NIGMS),
National Center for Research Resources under the PSI-2 Specialized
Center program (U54 GM074961); the National Institutes of Health
Roadmap for Medical Research (R01 GM075827 and P01
GM75913), and University of Chicago/Argonne National Laboratory
(ANL) Collaborative Seed Funding. We thank Nina Ponomarenko
and James R. Norris at the University of Chicago for samples of
Reaction Center from B. viridis. We thank Ray C. Stevens and Peter
Kuhn for helpful discussion and Elizabeth B. Haney for contributions
in writing and editing this manuscript. Use of the ANL General
Medicine and Cancer Institute Collaborative Access Team (GM/CA
CAT) beamlines at the Advanced Photon Source was supported by the
U.S. Department of Energy, Basic Energy Sciences, Office of Sci-
ence, under Contract No. DE-AC02-06CH11357. GM/CA CAT has
been funded in whole or in part with Federal funds from the National
Cancer Institute (Y1-CO-1020) and the NIGMS (Y1-GM-1104).
References
Ai X, Caffrey M (2000) Membrane protein crystallization in lipidic
mesophases: detergent effects. Biophys J 79(1):394–405
Caffrey M (2000) A lipid’s eye view of membrane protein crystal-
lization in mesophases. Curr Opin Struct Biol 10(4):486–497
Cherezov V, Caffrey M (2006) Picolitre-scale crystallization of
membrane proteins. J Appl Crystallogr 39:604–606
Cherezov V, Fersi H, Caffrey M (2001) Crystallization screens:
compatibility with the lipidic cubic phase for in meso crystal-
lization of membrane proteins. Biophys J 81(1):225–242
Cherezov V, Peddi A, Muthusubramaniam L, Zheng YF, Caffrey M
(2004) A robotic system for crystallizing membrane and soluble
proteins in lipidic mesophases. Acta Crystallogr D 60:1795–
1807
Cherezov V, Clogston J, Papiz MZ, Caffrey M (2006) Room to move:
crystallizing membrane proteins in swollen lipidic mesophases.
J Mol Biol 357(5):1605–1618
Cherezov V, Rosenbaum DM, Hanson MA, Rasmussen SGF, Thian
FS, Kobilka TS, Choi HJ, Kuhn P, Weis WI, Kobilka BK,
Stevens RC (2007) High-resolution crystal structure of an
engineered human beta(2)-adrenergic G protein-coupled recep-
tor. Science 318(5854):1258–1265
Cline SW, Doolittle WF (1987) Efficient transfection of the
archaebacterium Halobacterium halobium. J Bacteriol 169(3):
1341–1344
Davis J, Donohue TJ, Kaplan S (1988) Construction, characterization,
and complementation of a Puf- mutant of Rhodobacter sphaero-
ides. J Bacteriol 170(1):320–329
Duffy DC, McDonald JC, Schueller OJA, Whitesides GM (1998)
Rapid prototyping of microfluidic systems in poly(dimethylsi-
loxane). Anal Chem 70(23):4974–4984
Jaakola VP, Griffith MT, Hanson MA, Cherezov V, Chien EYT, Lane
JR, Ijzerman AP, Stevens RC (2008) The 2.6 angstrom crystal
structure of a human A(2A) adenosine receptor bound to an
antagonist. Science 322(5905):1211–1217
Katona G, Andreasson U, Landau EM, Andreasson LE, Neutze R
(2003) Lipidic cubic phase crystal structure of the photosynthetic
reaction centre from Rhodobacter sphaeroides at 2.35 angstrom
resolution. J Mol Biol 331(3):681–692
Lau BTC, Baitz CA, Dong XP, Hansen CL (2007) A complete
microfluidic screening platform for rational protein crystalliza-
tion. J Am Chem Soc 129(3):454–455
Li L, Mustafi D, Fu Q, Tereshko V, Chen DLL, Tice JD, Ismagilov
RF (2006) Nanoliter microfluidic hybrid method for simulta-
neous screening and optimization validated with crystallization
of membrane proteins. Proc Natl Acad Sci USA 103(51):19243–
19248
McDonald JC, Duffy DC, Anderson JR, Chiu DT, Wu HK, Schueller
OJA, Whitesides GM (2000) Fabrication of microfluidic systems
in poly(dimethylsiloxane). Electrophoresis 21(1):27–40
Nollert P (2002) From test tube to plate: a simple procedure for the
rapid preparation of microcrystallization experiments using the
cubic phase method. J Appl Crystallogr 35:637–640
Nollert P (2004) Lipidic cubic phases as matrices for membrane
protein crystallization. Methods 34(3):348–353
Ostermeier C, Michel H (1997) Crystallization of membrane proteins.
Curr Opin Struct Biol 7(5):697–701
Otwinowski Z, Minor W (1997) Processing of X-ray diffraction data
collected in oscillation mode. In: Carter CW, Sweet RM (eds)
Macromolecular crystallography, Pt A, vol 276, Academic Press
Inc., San Diego, pp 307–326
Peddi A, Muthusubramaniam L, Zheng YR, Cherezov V, Misquitta Y,
Caffrey M (2007) High-throughput automated system for
crystallizing membrane proteins in lipidic mesophases. IEEE
Trans Autom Sci Eng 4(2):129–140
Perry SL, Roberts GW, Tice JD, Gennis RB, Kenis PJA (2009)
Microfluidic generation of lipidic mesophases for membrane
protein crystallization. Cryst Growth Des 9(6):2566–2569
Microfluid Nanofluid
123

Pokkuluri PR, Laible PD, Deng YL, Wong TN, Hanson DK, Schiffer
M (2002) The structure of a mutant photosynthetic reaction
center shows unexpected changes in main chain orientations and
quinone position. Biochemistry 41(19):5998–6007
Roach LS, Song H, Ismagilov RF (2005) Controlling nonspecific
protein adsorption in a plug-based microfluidic system by
controlling interfacial chemistry using fluorous-phase surfac-
tants. Anal Chem 77(3):785–796
Shestopalov I, Tice JD, Ismagilov RF (2004) Multi-step synthesis of
nanoparticles performed on millisecond time scale in a micro-
fluidic droplet-based system. Lab Chip 4(4):316–321
Shim JU, Cristobal G, Link DR, Thorsen T, Jia YW, Piattelli K,
Fraden S (2007) Control and measurement of the phase behavior
of aqueous solutions using microfluidics. J Am Chem Soc
129(28):8825–8835
Simon R, Priefer U, Puhler A (1983) A broad host range mobilization
system for invivo genetic-engineering: transposon mutagenesis
in gram-negative bacteria. Bio-Technology 1(9):784–791
Song H, Tice JD, Ismagilov RF (2003) A microfluidic system for
controlling reaction networks in time. Angew Chem Int Ed
42(7):768–772
Song H, Chen DL, Ismagilov RF (2006a) Reactions in droplets in
microflulidic channels. Angew Chem Int Ed 45(44):7336–7356
Song H, Li HW, Munson MS, Van Ha TG, Ismagilov RF (2006b) On-
chip titration of an anticoagulant argatroban and determination
of the clotting time within whole blood or plasma using a plug-
based microfluidic system. Anal Chem 78(14):4839–4849
Taguchi AK W, Stocker JW, Alden RG, Causgrove TP, Peloquin JM,
Boxer SG, Woodbury NW (1992) Biochemical-characterization
and electron-transfer reactions of Sym1, a Rhodobacter capsul-
atus reaction center symmetry mutant which affects the initial
electron-donor. Biochemistry 31(42):10345–10355
Theiler R, Suter F, Zuber H, Cogdell RJ (1984) A comparison of the
primary structures of the 2 B800–850-apoproteins from wild-
type Rhodopseudomonas sphaeroides strain 2.4.1 and a carot-
enoidless mutant strain R26.1. FEBS Lett 175(2):231–237
Tiede DM, Vazquez J, Cordova J, Marone PA (1996) Time-resolved
electrochromism associated with the formation of quinone
anions in the Rhodobacter sphaeroides R26 reaction center.
Biochemistry 35(33):10763–10775
Toepke MW, Beebe DJ (2006) PDMS absorption of small molecules
and consequences in microfluidic applications. Lab Chip
6(12):1484–1486
Wadsten P, Wohri AB, Snijder A, Katona G, Gardiner AT, Cogdell
RJ, Neutze R, Engstrom S (2006) Lipidic sponge phase
crystallization of membrane proteins. J Mol Biol 364(1):
44–53
Wohri AB, Johansson LC, Wadsten-Hindrichsen P, Wahlgren WY,
Fischer G, Horsefield R, Katona G, Nyblom M, Oberg F, Young
G, Cogdell RJ, Fraser NJ, Engstrom S, Neutze R (2008) A
lipidic-sponge phase screen for membrane protein crystalliza-
tion. Structure 16(7):1003–1009
Zheng B, Roach LS, Ismagilov RF (2003) Screening of protein
crystallization conditions on a microfluidic chip using nanoliter-
size droplets. J Am Chem Soc 125(37):11170–11171
Zheng B, Gerdts CJ, Ismagilov RF (2005) Using nanoliter plugs in
microfluidics to facilitate and understand protein crystallization.
Curr Opin Struct Biol 15(5):548–555
Microfluid Nanofluid
123