Chemical processing strategies to obtain sporopollenin exine capsules from multi-compartmental pine pollen

Abstract

Pine pollen is widely used in traditional Chinese medicine and has been consumed as a food product for thousands of years. Owing to wind pollination, its pollen grains are composed of a sporoplasmic central cavity along with two empty air sac compartments. While this architectural configuration is evolutionarily optimized for wind dispersal, such features also lend excellent potential for encapsulating materials, especially in the context of preparing sporopollenin exine capsules (SECs). Herein, we systematically evaluated one-pot acid processing methods in order to generate pine pollen SECs that support compound loading. Morphological properties of the SECs were analysed by scanning electron microscopy (SEM) and dynamic imaging particle analysis (DIPA), and protein removal was evaluated by CHN elemental analysis and confocal laser scanning microscopy (CLSM). It was identified that 5-h acidolysis with 85% w/v phosphoric acid at 70 °C yielded an optimal balance of high protein removal and preservation of microcapsule architecture, while other processing methods were also feasible with an additional enzymatic step. Importantly, the loading efficiency of the pine pollen SECs was three-times greater than that of natural pine pollen, highlighting their potential for microencapsulation. Taken together, our findings outline a successful strategy to prepare intact pine pollen SECs and demonstrate for the first time that SECs can be prepared from multi-compartmental pollen capsules, opening the door to streamlined processing approaches to utilize pine pollen microcapsules in industrial applications.

Full text

Chemical
processing
strategies
to
obtain
sporopollenin
exine
capsules
from
multi-compartmental
pine
pollen
Arun
Kumar
Prabhakar
a,b
,
Hui
Ying
Lai
a,b
,
Michael
G.
Potroz
a,b
,
Michael
K.
Corliss
a,b
,
Jae
Hyeon
Park
a,b
,
Raghavendra
C.
Mundargi
a,b
,
Daeho
Cho
d
,
Sa-Ik
Bang
e,
**,
Nam-Joon
Cho
a,b,c,
*
a
School
of
Materials
Science
and
Engineering,
Nanyang
Technological
University,
50
Nanyang
Avenue,
639798
Singapore,
Singapore
b
Centre
for
Biomimetic
Sensor
Science,
Nanyang
Technological
University,
50
Nanyang
Drive,
637553
Singapore,
Singapore
c
School
of
Chemical
and
Biomedical
Engineering,
Nanyang
Technological
University,
62
Nanyang
Drive,
Singapore
637459,
Singapore
d
Department
of
Cosmetic
Sciences,
Sookmyung
Women’s
University,
Yongsan-ku,
Seoul
140-742,
Republic
of
Korea
e
Department
of
Plastic
Surgery,
Samsung
Medical
Center,
Sungkyunkwan
University
School
of
Medicine,
50
Ilwon-dong,
Gangnam-gu,
Seoul
135-710,
Republic
of
Korea
A
R
T
I
C
L
E
I
N
F
O
Article
history:
Received
16
March
2017
Received
in
revised
form
5
May
2017
Accepted
9
May
2017
Available
online
16
May
2017
Keywords:
Pollen
grains
Sporopollenin
exine
capsules
Sporopollenin
Pine
pollen
Microencapsulation
A
B
S
T
R
A
C
T
Pine
pollen
is
widely
used
in
traditional
Chinese
medicine
and
has
been
consumed
as
a
food
product
for
thousands
of
years.
Owing
to
wind
pollination,
its
pollen
grains
are
composed
of
a
sporoplasmic
central
cavity
along
with
two
empty
air
sac
compartments.
While
this
architectural
configuration
is
evolutionarily
optimized
for
wind
dispersal,
such
features
also
lend
excellent
potential
for
encapsulating
materials,
especially
in
the
context
of
preparing
sporopollenin
exine
capsules
(SECs).
Herein,
we
systematically
evaluated
one-pot
acid
processing
methods
in
order
to
generate
pine
pollen
SECs
that
support
compound
loading.
Morphological
properties
of
the
SECs
were
analysed
by
scanning
electron
microscopy
(SEM)
and
dynamic
imaging
particle
analysis
(DIPA),
and
protein
removal
was
evaluated
by
CHN
elemental
analysis
and
confocal
laser
scanning
microscopy
(CLSM).
It
was
identified
that
5-h
acidolysis
with
85%
w/v
phosphoric
acid
at
70

C
yielded
an
optimal
balance
of
high
protein
removal
and
preservation
of
microcapsule
architecture,
while
other
processing
methods
were
also
feasible
with
an
additional
enzymatic
step.
Importantly,
the
loading
efficiency
of
the
pine
pollen
SECs
was
three-times
greater
than
that
of
natural
pine
pollen,
highlighting
their
potential
for
microencapsulation.
Taken
together,
our
findings
outline
a
successful
strategy
to
prepare
intact
pine
pollen
SECs
and
demonstrate
for
the
first
time
that
SECs
can
be
prepared
from
multi-compartmental
pollen
capsules,
opening
the
door
to
streamlined
processing
approaches
to
utilize
pine
pollen
microcapsules
in
industrial
applications.
©
2017
The
Korean
Society
of
Industrial
and
Engineering
Chemistry.
Published
by
Elsevier
B.V.
All
rights
reserved.
Introduction
There
is
broad
interest
in
utilizing
natural
resources
to
develop
functional
materials
for
a
wide
range
of
applications
in
chemistry
and
materials
science
[1–4].
Given
their
diverse
range
of
highly
controlled
sizes
and
structures
that
vary
according
to
the
plant
species,
pollen
grains
as
well
as
related
spores
are
a
promising
example
of
a
natural
resource
that
offers
many
robust
options
for
microencapsulation
strategies
[5–7].
They
possess
a
large
cavity
surrounded
by
an
inner
structural
support
layer
(the
intine)
that
is
composed
of
cellulose,
hemicellulose,
and
pectin,
which
is
itself
surrounded
by
a
rigid
outer
coating
(the
exine)
that
is
composed
mainly
of
a
biopolymer,
sporopollenin
[8–10].
As
a
key
structural
component
that
supports
pollen
grains’
natural
function
to
protect
genetic
material,
sporopollenin
possesses
high
physical
and
chemical
resistance,
ultraviolet
shielding,
and
antioxidant
capa-
bility
[9,11–14].
Owing
to
these
features,
one
of
the
most
promising
directions
to
utilize
pollen
grains
for
microencapsulation
involves
the
chemical
processing
of
pollen
grains
to
yield
sporopollenin
exine
capsules
(SECs),
which
faithfully
preserve
the
architectural
features
of
the
original
grains
while
lacking
sporoplasmic
and
intine
contents
[15–25].
The
resulting
SECs
are
largely
devoid
of
*
Corresponding
author
at:
School
of
Materials
Science
and
Engineering,
Nanyang
Technological
University,
50
Nanyang
Avenue,
639798
Singapore,
Singapore.
**
Corresponding
author.
E-mail
addresses:
si55.bang@samsung.com
(S.-I.
Bang),
njcho@ntu.edu.sg
(N.-J.
Cho).
http://dx.doi.org/10.1016/j.jiec.2017.05.009
1226-086X/©
2017
The
Korean
Society
of
Industrial
and
Engineering
Chemistry.
Published
by
Elsevier
B.V.
All
rights
reserved.
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
Contents
lists
available
at
ScienceDirect
Journal
of
Industrial
and
Engineering
Chemistry
journal
homepa
ge:
www.elsev
ier.com/locate/jie
c

protein,
making
them
less
allergenic
and
offering
more
volume
for
molecular
encapsulation
[26].
Combined
with
high
natural
abundance
and
renewable
production,
these
advantageous
prop-
erties
make
SECs
an
ideal
delivery
vehicle
for
encapsulating
therapeutic
drugs,
nutrients,
and
microorganisms
as
well
as
for
other
material
science
applications
[26–36].
Pollen
processing
has
mainly
focused
on
single-compartment
pollen
species,
typically
ones
which
function
via
biotic
pollination
and
hence
possess
thicker
exine
walls
[37,38].
Early
attempts
to
prepare
SECs
involved
sequential
treatments
of
organic
solvents,
alkali,
and
acid
in
order
to
remove
lipids,
proteins,
and
intine
layers
[39–42].
One-pot
methods
aimed
at
streamlining
the
process
were
later
introduced
and
included
individual
treatments
with
hydro-
chloric
acid
[15,16],
acetic
acid
[19,21],
hydrofluoric
acid
[18],
and
combinations
of
aqueous
4-methylmorpholine-N-oxide
and
su-
crose
[17,20].
Multi-compartmental
pollens,
however,
including
saccate
pollens
that
travel
by
wind
dispersal,
are
generally
more
fragile
due
to
thinner
exine
walls
and
often
rupture
during
harsh
chemical
treatments.
Indeed,
these
challenges
have
proven
a
common
theme
among
thin-walled
pollen,
and
have
motivated
the
search
for
improved
processing
strategies
[43].
As
a
result,
milder
processing
methods
such
as
purely
enzymatic
treatments
have
been
developed
[39–42].
However,
enzymatic
processing
is
not
practically
feasible
at
larger
scales
because
the
necessary
enzymes
are
costly
and
required
in
relatively
high
concentrations.
Develop-
ing
robust
chemical
methods
to
prepare
SECs
from
multi-
compartmental
pollen
grains
would
increase
the
general
utility
of
pollen
SECs,
especially
since
these
grains
are
among
the
most
abundant
in
nature
[38].
In
this
regard,
pine
(Pinus
taeda)
pollen
is
noteworthy
because
it
has
been
widely
used
in
traditional
Chinese
medicine
for
thousands
of
years
[44]
and
its
structure
has
been
rigorously
studied
in
the
biological
sciences
[45–48].
Pine
pollen
has
two
lightweight,
empty
air
sacs
attached
to
its
sporoplasmic
central
cavity
that
aid
in
wind
pollination
and
the
compartments
collectively
form
a
micron-scale
size
(45–75
mm)
tripartite
structure
[49,50].
These
relatively
fragile
sacci
possess
higher
permeability
on
account
of
porous
features
that
are
nearly
two
orders
of
magnitude
larger
than
those
of
the
central
cavity
(200
nm
vs
4
nm)
[51],
and
understanding
how
these
different
permeability
barriers
might
be
useful
for
microencapsulation
remains
to
be
explored
[5,52,53].
Indeed,
while
the
membrane
permeability
properties
of
isolated
pine
pollen
exines
have
been
investigated
in
the
context
of
fundamental
pollen
biology
[50,51,54],
there
have
been
no
attempts
to
develop
focused
strategies
for
preparing
pine
pollen
SECs.
Addressing
this
gap
would
significantly
advance
efforts
to
achieve
scalable
chemical
processing
methods
for
obtaining
high-quality
pine
pollen
SECs
as
well
as
to
explore
the
feasibility
of
loading
the
SECs
with
macromolecules
for
microen-
capsulation
applications.
Indeed,
while
pine
pollen
has
been
shown
to
have
relatively
low
allergenicity
(see,
e.g.,
its
low
skin
sensitivity
[55,56]
and
steric
limitations
for
pulmonary
uptake
[57]),
SEC
development
could
further
improve
this
species’
immunogenic
profile
[58]
as
well
as
increase
the
scope
of
applications
based
on
enhanced
loading
of
selected
compounds
into
hollow
cavities.
Herein,
we
conducted
the
first
systematic
evaluation
of
one-pot
acidolysis
protocols
for
pine
pollen
SEC
extraction
in
addition
to
comprehensive
characterization
of
pine
pollen
SEC
morphology
and
encapsulation
efficiency.
Processing
times,
storage
methods,
temperatures,
and
acid
type
were
varied
in
order
to
achieve
maximum
removal
of
potentially
allergenic
protein
components
while
preserving
microcapsule
architecture.
In
selected
cases,
additional
enzymatic
treatment
with
trypsin
was
used
to
further
demonstrate
the
potential
of
fully
optimizing
sporoplasmic
removal.
A
comprehensive
set
of
experimental
techniques
was
utilized
in
order
to
characterize
the
morphological
and
chemical
properties
of
processed
samples
as
well
as
to
investigate
the
loading
of
macromolecules.
Experimental
Materials
Defatted
P.
taeda
pollen
was
purchased
from
Greer
Laboratories
(Lenoir,
NC,
USA).
Sodium
bicarbonate,
sodium
dodecyl
sulphate,
sodium
chloride,
organic
solvents
(acetone,
ethanol),
BSA,
and
FITC-BSA
were
obtained
from
Sigma–Aldrich
(Singapore).
Phos-
phoric
acid
(85%
w/v)
was
procured
from
Merck
(Singapore).
Hydrochloric
acid
(37%
w/v)
and
sulphuric
acid
(95%
w/v)
were
purchased
from
VWR
Chemical
(Singapore).
Polystyrene
micro-
spheres
(50

1
mm
diameter)
and
0.25%
Trypsin-EDTA
were
purchased
from
Thermo
Fischer
Scientific
(Waltham,
MA,
USA).
Extraction
of
pine
sporopollenin
exine
capsules
(SECs)
Acidolysis
processing
Defatted
pine
pollen
(2
g)
was
placed
into
a
round-bottomed
perfluoroalkoxy
(PFA)
flask
fitted
with
a
glass
condenser
and
refluxed
(70

C)
in
aqueous
85%
phosphoric
acid
solution
(15
mL)
at
a
stirring
rate
of
200
rpm
for
one
hour.
Samples
were
collected
by
vacuum
filtration
and
washed
sequentially
with
warm
distilled
water
(150
mL

5),
acetone
(100
mL

1),
2
M
HCl
(100
mL

1),
distilled
water
(100
mL

5),
acetone
(100
mL

1),
and
ethanol
(100
mL

2).
Washed
microcapsules
were
dried
in
an
oven
at
60

C
for
6
h,
followed
by
storage
at
room
temperature
as
either
a
dry
powder
or
aqueous
suspension
in
water.
This
protocol
was
repeated
for
different
batches
of
the
same
pollen,
using
different
reflux
durations
(1
h,
2.5
h,
5
h,
10
h,
and
20
h)
and
samples
were
collected
for
analytical
characterization.
The
reflux
duration
that
demonstrated
the
highest
yield
of
intact
pollen
microcapsules
was
re-tested
using
one
or
more
of
the
following
adjustments:
wet
storage,
as
a
suspension
of
SECs
in
distilled
water
that
is
kept
at
room
temperature;
lowered
reflux
temperatures
(50

C
and
25

C);
and
different
acid
reagents
(hydrochloric
and
sulphuric
acids).
Enzymatic
treatment
with
trypsin
As
an
extension
of
the
study,
one
sample
(18%
w/v
hydrochloric
acid
for
5
h
at
70

C)
was
chosen
for
additional
enzymatic
treatment
with
trypsin.
This
batch
was
washed
with
room
temperature
distilled
water
(100
mL

5)
over
vacuum
filtration
and
transferred
into
a
0.25%
trypsin-EDTA
(15
mL)
solution
for
24
h
incubation
at
35

C.
Samples
were
subsequently
washed
with
distilled
water
(100
mL

5),
transferred
into
sodium
bicarbonate
solution
(10
g/L)
containing
sodium
dodecyl
sulphate
(1
g/L),
and
finally
dried
for
24
h
at
room
temperature.
After
drying,
the
SECs
were
thoroughly
washed
with
distilled
water
and
divided
into
wet
and
dry
storage.
Micromeritic
evaluation
by
dynamic
imaging
particle
analysis
(DIPA)
DIPA
was
performed
using
a
FlowCam
1
benchtop
system
(FlowCamVS,
Fluid
Imaging
Technologies,
Maine,
USA)
equipped
with
a
200
mm
width
flow
cell
(FC-200)
and
20
magnification
lens
(Olympus
1
,
Japan).
The
flow
cell
was
visually
inspected
and
cleaned
with
ethanol
prior
to
each
sample
run.
Polystyrene
microspheres
(50

1
mm
diameter)
served
to
calibrate
the
microscope
focus,
and
a
representative
histogram
was
plotted
to
verify
the
proper
operation
under
the
defined
measurement
settings.
Defatted
natural
pine
pollen
and
processed
SECs
(at
2
mg/
mL
concentration)
were
sonicated
in
a
water
bath
for
10
min
and
filtered
through
100
mm
diameter
filter
meshes
prior
to
experi-
ment.
The
samples
were
then
manually
added
into
the
flow
cell
via
376
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385

a
pump-controlled
syringe
and
analysed
at
a
flow
rate
of
0.1
mL/
min.
A
minimum
of
10,000
particles
was
scanned
and
three
separate
measurements
were
performed.
Data
analysis
was
carried
out
using
300
well-focused
particles.
Surface
morphology
evaluation
by
scanning
electron
microscopy
(SEM)
SEM
imaging
was
performed
using
a
JSM
5410
(JEOL,
Tokyo,
Japan).
Samples
were
sputter-coated
with
a
10
nm-thick
gold
film
using
a
JFC-1600
instrument
(JEOL,
Tokyo,
Japan)
at
20
mA
for
60
s.
Images
were
captured
at
an
accelerating
voltage
of
5
kV
at
different
magnifications
and
both
interior
(cross-sectional)
and
exterior
morphological
changes
were
observed.
Elemental
CHN
analysis
A
VarioEL
III
elemental
analyser
(Elementar,
Hanau,
Germany)
provided
CHN
analysis
to
determine
the
amount
of
residual
protein.
All
samples
were
dried
at
60

C
for
1
h
before
being
combusted
in
excess
oxygen
at
high
temperature
to
release
compositional
carbon,
hydrogen,
and
nitrogen.
Protein
content
was
calculated
using
percent
nitrogen
with
a
6.25
multiplication-
factor
that
corresponds
to
the
protein
content
in
pine
pollen
SECs,
in
accordance
to
recommendations
from
the
Association
of
Analytical
Communities
[59].
All
measurements
were
conducted
in
triplicate.
Confocal
laser
scanning
microscopy
analysis
(CLSM)
Defatted
pine
pollen,
processed
SECs,
and
fluorescein
isothio-
cyanate-bovine
serum
albumin
(FITC-BSA)-loaded
SECs
were
mounted
on
sticky
slides
with
Vectashield
1
and
scanned
via
confocal
laser
scanning
microscopy
(Carl
Zeiss
LSM700,
Germany),
as
previously
described
[60].
Laser
excitation
lines
were
set
to
405
nm,
488
nm,
and
561
nm
at
a
scan
speed
of
67
s
per
phase.
Images
were
collected
with
differential
interference
contrast
at
405
nm
(6.5%),
488
nm
(6%)
and
561
nm
(6%)
using
enhanced-
contrast
(EC)
Plan-Neofluar
100 
and
150 0

1.3
oil
objective
M27
lenses.
The
fluorescence
emission
was
collected
in
photomultiplier
tubes
equipped
with
different
filters
(416–477
nm,
498–5
nm,
and
572–620
nm)
and
analysed
by
using
the
ZESS
2008
software
package.
Encapsulation
of
bovine
serum
albumin
(BSA)
BSA
protein
loading
was
achieved
by
using
vacuum
loading
protocols
[54,61].
Unprocessed
pollen
and
select
batches
of
dry-
stored
processed
SECs
(85%
phosphoric
acid
for
5
h
at
70

C,
and
18%
hydrochloric
acid
for
5
h
at
70

C,
with
additional
24
h
trypsin
treatment)
were
suspended
in
0.5
mL
of
50
mg/mL
aqueous
BSA
solution
within
polypropylene
tubes
and
mixed
via
vortexing
(IKA,
Staufen,
Germany)
for
1
min.
The
mixture
was
then
subjected
to
vacuum
treatment
at
0.006
mbar
for
2
h.
The
tubes
were
collected
and
the
loaded
particles
were
washed
by
centrifugation
with
1
mL
of
water
at
12,000
rpm
for
3
min
and
then
freeze-dried
overnight.
Blank
batches
of
untreated
pollen
and
SECs
were
prepared
similarly
without
BSA
loading.
In
addition,
FITC-BSA
was
encapsu-
lated
in
the
same
way
and
imaged
via
CLSM
in
order
to
observe
the
molecular
localization
of
loaded
components
within
SEC
particles.
Following
preparation,
BSA-loaded
SECs
(10
mg)
were
crushed
using
a
mortar
and
pestle
for
5
min
to
expel
encapsulated
BSA
[62].
The
crushed
powder
was
mixed
with
phosphate-buffer
saline
[pH
7.4]
(2
mL),
vortexed
for
5
min,
and
centrifuged
at
15,000
rpm
for
5
min.
The
supernatant
was
filtered
using
a
0.45
mm
diameter
polyethersulfone
(PES)
syringe
filter
(Agilent,
CA,
USA).
Absorbance
values
were
measured
at
280
nm
by
using
a
Boeco-
S220
spectrophotometer
(Hamburg,
Germany)
along
with
appro-
priate
controls
(SEC
blank
and
BSA
standards
at
different
concentrations),
and
the
amount
of
BSA
in
the
SECs
was
calculated
by
the
following
equation:
%
BSA
loading
¼Weight
of
BSA
Weight
of
BSA

loaded
SECs 
100%
Results
and
discussion
Process
development
and
analytical
characterization
Phosphoric
acid
(H
3
PO
4
)
is
widely
used
in
consumer
products,
generally-recognized-as-safe,
and
affordable,
and
has
consequent-
ly
proven
an
attractive
solvent
for
SEC
processing
[11,22,59].
Pollen
SEC
extraction
using
phosphoric
acidolysis
has
been
convention-
ally
performed
at
temperatures
up
to
180

C
for
durations
as
long
as
one
week
(168
h)
[26,32–35,52,63].
While
such
protocols
may
suit
SEC
extraction
from
single-compartment,
thick-walled
pollen
grains
and
spores
such
as
Lycopodium
clavatum,
they
can
cause
significant
damage
to
the
tripartite
microstructure
of
P.
taeda
pollen
grains,
including
the
fragile
air
sacs
[51].
In
recent
work,
our
group
has
successfully
established
streamlined
acidolysis
proto-
cols
to
extract
SECs
from
Healianthus
annuus
(sunflower)
[64],
L.
clavatum
(moss)
[61]
and
Zea
mays
(corn)
[43]
using
highly
efficient
acidolysis
protocols
that
require
much
shorter
time
intervals.
Building
on
these
efforts
with
single-compartment
pollen
grains
and
in
light
of
the
challenges
associated
with
multi-compartment
pine
pollen,
the
following
experimental
and
analytical
characterization
strategies
were
aimed
at
identifying
suitable
processing
strategies
to
extract
pine
pollen
SECs.
Processing
scheme
As
presented
in
Fig.
1,
we
systematically
tested
one-pot
phosphoric
acid
processing
across
a
variety
of
durations,
temper-
atures,
and
storage
methods
based
on
the
following
SEC
extraction
process:
natural
pine
pollen
was
first
defatted
with
diethyl
ether,
refluxed
with
acid
under
the
appropriate
conditions,
and
then
washed
and
dried.
Extracted
pine
SECs
were
stored
in
either
dry
(conventional)
or
aqueous
wet
conditions
(to
mitigate
potential
collapse
of
the
thin-walled
pollen),
and
characterized
using
various
analytical
techniques
in
order
to
evaluate
the
degree
of
structural
preservation
as
well
as
the
removal
of
sporoplasmic
and
protein
contents.
From
these
experiments,
optimal
conditions
were
identified
and
then
extended
to
strong
acids
and,
in
some
cases,
the
treatment
protocols
included
an
additional
enzymatic
processing
step
[15,16].
A
detailed
description
of
all
processing
conditions
is
provided
in
Table
1.
Evaluation
of
SEC
structural
preservation
Depending
on
the
extraction
process,
the
structural
integrity
of
the
resulting
SECs
will
vary
and
the
morphological
structure
of
SEC
particles
can
be
analysed
at
the
single-particle
level
by
high-
throughput
DIPA
measurements
[61].
We
divided
the
pine
pollen
SECs
into
three
groups
for
classification:
“intact,”
“fractured,”
and
“collapsed”.
Representative
examples
of
particles
that
were
assigned
to
each
group
are
presented
in
Fig.
2.
Preserved
particles
closely
resemble
unprocessed
pine
pollen,
with
both
air
sacs
attached
to
the
central
cavity
in
a
tripartite
microstructure
and
show
no
significant
breaks
or
cracks.
However,
they
may
have
minor
deviations
in
shape,
average
diameter,
or
increases
in
surface
roughness,
as
compared
to
the
untreated
case.
Fractured
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
377

SECs
have
clearly
visible
holes,
cracks,
or
missing
sections
in
one
or
more
compartment,
but
at
least
one
uncompromised,
fully
enclosed
compartment
remaining
based
on
the
visual
inspection.
Examples
of
fractured
SECs
may
include
a
complete
central
cavity
that
has
damaged
or
missing
air
sacs,
or
an
intact
air
sac
that
is
attached
to
other
damaged/missing
compartments.
Collapsed
SECs
appear
shrivelled
and
flat
and
have
very
little
inner
volume
for
loading.
These
classifications
provide
a
starting
point
to
aid
the
development
of
optimized
processing
strategies.
Effects
of
processing
conditions
on
particle
intactness
Following
this
characterization
strategy,
we
initially
examined
the
effects
of
processing
duration,
storage
method,
and
reflux
temperature
on
SEC
particle
morphology,
and
the
results
are
summarized
in
Table
2.
Before
processing,
the
natural
pollen
grains
were
99%
intact
with
only
trace
amounts
of
collapsed
and
fractured
particles.
The
first
parameter
that
was
then
tested
was
the
time
scale
of
pollen
grain
processing
in
85%
phosphoric
acid
at
70

C,
followed
by
conventional
dry
storage.
After
1
h
of
processing,
45%
of
particles
were
intact
while
32%
were
collapsed.
With
increasing
processing
time,
the
percentage
of
intact
particles
increased
to
60%
while
the
percentage
of
collapsed
particles
decreased
to
approximately
10%
and
2%
after
5
h
and
20
h,
respectively.
This
decrease
in
the
fraction
of
collapsed
particles
at
longer
processing
times
supports
past
accounts
of
pine
pollen
grains
undergoing
rapid
collapse
due
to
acid
shock
resulting
from
phosphate
ion
exchange
between
membranes
[32].
At
the
same
time,
processing
times
of
10
h
or
longer
led
to
a
large
number
of
fractured
particles,
reaching
around
40%.
Optimal
results
in
this
test
series
were
obtained
with
processing
in
85%
phosphoric
acid
at
70

C
for
5
h,
with
60%
intact
particles,
as
well
as
30%
fractured
and
10%
collapsed.
As
the
sporopollenin
exine
walls
of
pine
pollen
are
relatively
thin
and
hence
fragile,
we
next
explored
whether
a
wet
storage
environment
that
provides
structural
stability
[43]
could
reduce
the
percentage
of
collapsed
particles
and
improve
the
overall
yield.
To
explore
this
option,
the
particles
were
processed
in
85%
phosphoric
acid
at
70

C,
followed
by
wet
storage.
In
this
case,
the
percentage
of
intact
particles
increased
to
78%
while
the
percentage
of
collapsed
particles
decreased
to
0.5%.
These
findings
support
that
wet
storage
facilitates
particle
intactness
by
avoiding
the
structural
collapse/buckling
that
occurs
with
dehydrated
pollen
microcapsules
[65–68],
although
dry
storage
would
likely
increase
durability
and
shelf-life
of
the
SEC
particles
for
industrial
applications.
Similar
results
were
also
obtained
with
processing
in
42%
phosphoric
acid,
supporting
that
the
high
acid
concentration
is
suitable
for
SEC
production.
Based
on
this
optimized
condition
(5
h
phosphoric
acid,
wet
storage),
the
effect
of
temperature
was
next
investigated
and
the
number
of
intact
particles
increased
at
lower
temperature,
reaching
93%
intact
particles
when
processed
at
25

C.
Hence,
good
control
over
the
processing
steps
could
be
achieved
as
indicated
by
retention
of
pine
pollen
exine
morpholo-
gy
in
the
SECs.
For
comparison,
we
used
a
similar
standard
protocol
(5
h
at
70

C)
to
test
two
strong
acids,
including
hydrochloric
(HCl)
and
Table
1
Processing
parameters
used
for
extraction
of
Pinus
taeda
sporopollenin
exine
capsules.
Conditions
Processing
parameters
Reagent
Study
Time
(h)
Conc.
(w/v)
Storage
Temp.
(

C)
Unprocessed
–
–
–
–
–
H
3
PO
4
Time
1
85%
Dry
70
2.5
85%
Dry
70
5
85%
Dry
70
10
85%
Dry
70
20
85%
Dry
70
Storage
5
85%
Wet
70
5
42%
Wet
70
Temperature
5
85%
Wet
50
5
85%
Wet
25
HCl
Strong
acid
5
18%
Wet
70
5
27%
Wet
70
H
2
SO
4
5
25%
Wet
70
Fig.
1.
Chemical
processing
strategy
to
extract
pine
pollen
sporopollenin
exine
capsules
(SECs).
Defatted
Pinus
taeda
pollen
grains,
dispersed
by
wind,
are
collected
in
the
natural
state
and
prepared
by
incubation
in
diethyl
ether
to
remove
lipid
components.
Then,
the
grains
are
subjected
to
acidolysis
in
a
specific
acidic
solvent
(strong
or
weak
acid)
and
incubation
conditions
to
remove
proteinaceous
content.
Sequential
washings
are
performed
to
remove
residual
solvent
from
processed
capsules,
and
the
capsules
are
subjected
to
analytical
characterization
for
quality
control,
including
morphological
assessment
and
protein
removal
efficiency.
378
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385

sulphuric
(H
2
SO
4
)
acids.
Processing
in
18%
hydrochloric
acid
successfully
yielded
92%
intact
and
7%
fractured
particles,
while
processing
in
27%
hydrochloric
acid
was
less
optimal,
resulting
in
83%
intact
and
17%
fractured
particles.
On
the
other
hand,
processing
in
sulphuric
acid
yielded
77%
intact
and
23%
fractured
particles.
As
these
two
strong
acids
are
known
to
dissolve
cellulosic
intine
materials
but
not
proteinaceous
materials
[69],
we
also
explored
the
feasibility
of
adding
an
additional
processing
step
with
trypsin
protease
(as
described
in
Ref.
[51])
to
the
18%
hydrochloric
acid
protocol
and
identified
that
the
number
of
intact
particles
decreased
to
75%.
Collectively,
the
findings
demonstrate
that
both
weak
and
strong
acids
that
are
widely
used
in
SEC
extraction
protocols
successfully
work
with
pine
pollen
grains,
and
that
the
fragile
nature
of
this
thin-walled
pollen
species
is
an
important
factor
for
optimizing
the
processing
conditions.
Based
on
the
data
collected,
the
initial
evidence
suggests
that
a
large
percentage
of
intact
particles
can
be
obtained
with
single-pot
phosphoric
acid
processing
or
a
combination
of
strong
acid
(e.g.,
Fig.
2.
Representative
optical
micrographs
of
processed
SECs
with
different
morphological
states.
Based
on
visual
inspection
of
the
optical
micrographs,
individual
particles
were
classified
as
(A)
intact
(preserved
tripartite
microstructure
with
no
ostensible
breaks
or
cracks),
(B)
fractured
(cracked
or
missing
portions,
with
at
least
one
fully
preserved
compartment),
or
(C)
collapsed
(significantly
shrivelled
with
low
encapsulation
volume).
Table
2
Effect
of
different
processing
conditions
on
the
morphological
properties
of
processed
SEC
samples.
The
number
of
intact,
fractured,
and
collapsed
particles
are
expressed
as
percentages
from
data
collected
for
>300
particles.
Morphological
characterization
Processing
condition
Intact
(%)
Fractured
(%)
Collapsed
(%)
Unprocessed
99.0

0.3
0.8

0.2
0.1

0.2
85%
H
3
PO
4
at
70

C
for
1
h
(dry)
44.8

6.6
23.2

6.0
31.8

4.2
85%
H
3
PO
4
at
70

C
for
2.5
h
(dry)
56.6

5.8
24.1

2.4
18.6

2.6
85%
H
3
PO
4
at
70

C
for
5
h
(dry)
60.6

0.8
29.8

1.9
9.4

2.3
85%
H
3
PO
4
at
70

C
for
10
h
(dry)
55.2

8.7
41.2

9.2
3.7

0.50
85%
H
3
PO
4
at
70

C
for
20
h
(dry)
55.2

3.8
43.2

4.1
1.6

0.6
85%
H
3
PO
4
at
70

C
for
5
h
(wet)
80.6

4.2
18.7

4.3
0.5

0.00
42%
H
3
PO
4
at
70

C
for
5
h
(wet)
76.1

8.7
23.3

8.7
0.5

0.5
85%
H
3
PO
4
at
50

C
for
5
h
(wet)
82.0

3.6
18.0

3.6
0.0

0.0
85%
H
3
PO
4
at
25

C
for
5
h
(wet)
93.3

3.2
6.7

3.2
0.0

0.0
18%
HCl
at
50

C
for
5
h
(wet)
77.1

6.4
22.8

6.4
0.0

0.0
27%
HCl
at
25

C
for
5
h
(wet)
92.2

2.0
7.2

2.1
0.1

0.2
25%
H
2
SO
4
at
50

C
for
5
h
(wet)
83.0

3.7
17.0

3.7
0.0

0.0
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
379

hydrochloric
acid)
processing
followed
by
enzymatic
treatment
with
a
protease
enzyme.
Micromeritic
properties
In
addition
to
assessing
the
structural
integrity
of
chemically
processed
samples,
DIPA
analysis
was
conducted
in
order
to
determine
the
number-weighted
average
particle
diameter
of
each
sample,
as
presented
in
spline
curve
fit
histograms
in
Fig.
3.
While
unprocessed
pollen
grains
had
an
average
diameter
of
62
mm,
the
average
diameter
shrank
to
around
56
mm
for
samples
treated
with
85%
(70

C)
phosphoric
acidolysis
for
2.5
h
or
shorter
durations.
With
longer
processing
times,
the
average
particle
diameters
once
again
approached
the
values
for
unprocessed
pollen
grains
(Fig.
3A),
and
the
average
diameter
of
particles
processed
for
5
h
duration
or
longer
was
around
59
mm.
Furthermore,
the
average
diameters
of
particles
treated
with
85%
or
42%
phosphoric
acid
were
similar,
as
were
the
samples
processed
at
different
temper-
atures,
once
again
demonstrating
that
the
high
acid
concentration
and
high
temperature
are
suitable
for
the
processing
step
(Fig.
3B,
C).
The
strong
acid
treatments
also
did
not
affect
the
particle
size
(Fig.
3D).
Taken
together,
the
data
reinforce
that
an
optimal
processing
time
(in
the
range
of
5–10
h,
and
defined
to
be
5
h
for
our
experiments)
overcomes
initial
particle
collapse
due
to
acid
shock
while
avoiding
excessive
particle
damage
due
to
fracturing.
Morphological
investigation
Representative
SEM
micrographs
of
pine
pollen
grains
after
phosphoric
acid
acidolysis
for
different
processing
times
are
presented
in
Fig.
4.
The
cross-sectional
image
of
the
unprocessed
pine
pollen
grains
reveals
the
presence
of
pollen
constituents
inside
the
large
central
inner
cavity
(Fig.
4A).
As
discussed
above,
the
majority
of
SEC
particles
appeared
to
collapse
after
acidolysis
for
a
period
of
1
h
and
some
were
still
present
after
2.5
h
(Fig.
4B,
C).
However,
at
longer
processing
times,
the
fraction
of
collapsed
particles
decreased
and
the
5
h
time
point
again
showed
an
optimal
balance
of
intact
particles
(Fig.
4D–F).
In
addition,
the
SEM
micrographs
for
pine
pollen
SECs
prepared
using
strong
acids
are
presented
in
Fig.
5.
In
all
three
tested
cases,
the
SECs
appear
largely
intact
as
expected.
While
most
particles
were
observed
to
be
intact,
the
fraction
of
damaged
particles
were
generally
more
fractured
than
collapsed.
Indeed,
cross-section
images
were
obtained
from
highly
damaged
particles
and,
in
these
cases,
proteinaceous
components
were
still
visible
in
the
central
cavity.
These
aspects
are
further
discussed
below
in
the
context
of
elemental
analysis
and
highlight
that
phosphoric
acid
is
advantageous
for
removing
sporoplasmic
contents
in
general,
while
strong
acids
require
an
additional
processing
step
with
a
proteolytic
enzyme
to
aid
SEC
production.
Fig.
3.
Size
histograms
of
processed
SEC
particles.
(A)
Effect
of
processing
time
in
85%
phosphoric
acid
at
70

C.
(B)
Effect
of
phosphoric
acid
concentration.
The
time
and
temperature
were
fixed
at
5
h
and
at
70

C,
respectively.
(C)
Effect
of
processing
temperature
for
5
h
processing
in
85%
phosphoric
acid.
(D)
Alternative
processing
strategies
with
strong
acids
(5
h
at
70

C).
Data
is
collected
from
>300
individual
particles
per
sample.
380
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385

Fig.
4.
SEM
micrographs
of
(A)
unprocessed
pine
pollen
and
(B–F)
SECs
processed
with
85%
phosphoric
acid
at
70

C
for
varying
durations
of
processing
time.
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
381

Fig.
5.
SEM
micrographs
of
SECs
processed
with
different
strong
acids
for
5
h
at
70

C.
Fig.
6.
Protein
content
and
protein
removal
efficiency
for
processed
SEC
samples
as
determined
by
CHN
analysis.
The
removal
efficiency
is
determined
based
on
the
protein
content
of
the
processed
samples
relative
to
an
unprocessed
sample.
382
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385

Assessment
of
protein
removal
In
addition
to
retaining
morphological
structure,
a
key
requirement
of
SEC
production
is
the
removal
of
potentially
allergenic
proteins
from
the
pollen
microcapsules.
To
verify
protein
removal,
CHN
elemental
analysis
based
on
high-temperature
combustion
was
conducted
on
untreated
and
treated
pine
pollen
samples,
and
the
percentage
of
protein
removal
was
determined
based
on
the
amount
of
nitrogen
remaining
in
the
samples.
The
key
measurement
principle
behind
this
approach
is
that
it
is
known
that
proteins
are
the
only
major
component
of
pollen
grains
which
contain
nitrogen,
and
hence
measuring
the
nitrogen
content
of
SEC
particles
provides
an
indication
of
how
much
protein
was
removed
as
a
result
of
chemical
processing
[52].
The
percentage
of
sporoplasmic
protein
removal
was
determined
by
the
following
equation:
%Sporoplasmic
removal
¼
%N
Unprodessed

%N
Processed
%N
Unprocessed

100%
As
presented
in
Fig.
6,
unprocessed
pine
pollen
was
determined
to
have
a
protein
content
of
around
11%,
which
agrees
well
with
literature
values
for
other
pollen
species
tested
in
our
group
(8–32%
depending
on
species)
and
this
control
sample
provided
the
reference
value
for
0%
protein
removal.
While
1
h
treatment
with
85%
phosphoric
acid
at
70

C
led
to
a
70%
reduction
in
protein
content,
longer
treatments
with
85%
phosphoric
acid
were
more
effective,
yielding
around
85–89%
removal
of
protein
content.
This
efficiency
agrees
well
with
SECs
prepared
from
other
pollen
species.
When
using
85%
phosphoric
acid
at
70

C,
treatment
times
of
2.5
h
or
longer
were
equivalent
in
their
utility
for
protein
removal.
By
contrast,
treatment
with
85%
phosphoric
acid
at
lower
temperatures
was
less
effective
at
removing
proteins,
achieving
removal
efficiencies
around
75%.
Likewise,
5
h
treatment
with
42%
phosphoric
acid
at
70

C
also
had
poor
removal
efficiency,
around
63%.
Taken
together,
these
data
support
that
5
h
treatment
with
85%
phosphoric
acid
at
70

C
was
particularly
effective
at
removing
protein
and,
in
line
with
the
morphological
analysis,
this
processing
condition
was
selected
from
among
the
one-pot
options
as
the
optimal
strategy
for
preparing
SECs
for
microen-
capsulation.
On
the
other
hand,
treatments
with
strong
acids
were
less
effective
at
removing
protein.
In
all
cases,
one-pot
treatment
of
pine
pollen
grains
with
strong
acids
(5
h
treatment
at
70

C)
led
to
protein
removal
efficiencies
around
57–65%.
These
values
are
consistent
with
the
previous
observations
that
indicate
that
strong
acids
are
typically
less
effective
at
cleaning
SECs.
To
improve
protein
removal,
trypsin
was
added
to
the
hydrochloric
Fig.
7.
CLSM
images
of
unprocessed
pine
pollen
and
processed
SEC
samples
without
and
with
loaded
BSA.
The
processing
conditions
were
either
85%
phosphoric
acid
for
5
h
at
70

C,
or
18%
hydrochloric
acid
for
5
h
at
70

C
followed
by
trypsin
treatment.
Left
column:
cross-section
of
pollen
or
SECs
before
protein
loading.
The
blue
autofluorescence
corresponds
to
compounds
naturally
present
in
pollen
or
SECs.
The
other
columns
present
3D
reconstructions
of
BSA-loaded
pollen
or
SECs,
for
which
the
dual-channel
CLSM
images
show
compounds
naturally
present
in
pollen
or
SECs
(blue)
and
loaded
FITC-labeled
BSA
(green).
All
scale
bars
are
20
m
m.
(For
interpretation
of
the
references
to
color
in
this
figure
legend,
the
reader
is
referred
to
the
web
version
of
this
article.)
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
383

acid-treated
pollen
sample
and
this
two-step
protocol
had
a
protein
removal
efficiency
around
93%.
As
a
control,
it
was
also
identified
that
trypsin
alone
was
ineffective
for
removing
protein.
Hence,
sequential
treatments
with
hydrochloric
acid
and
trypsin
were
selected
as
the
two-step
method
of
choice
for
preparing
SECs
for
further
exploration
for
microencapsulation.
Evaluation
of
loading
efficiency
Fig.
7
presents
confocal
laser
scanning
microscopy
(CLSM)
images
characterizing
the
loading
properties
of
unprocessed
and
processed
pollen
grains.
As
discussed
above,
two
SEC
samples
were
selected
for
this
evaluation
based
on
one-pot
treatment
(85%
phosphoric
acid
for
5
h
at
70

C)
and
two-step
treatment
(18%
hydrochloric
acid
for
5
h
at
70

C,
followed
by
trypsin
incubation),
respectively.
The
CLSM
approach
is
useful
because
the
pollen
exine
wall
as
well
as
its
sporoplasmic
contents
are
known
to
autofluoresce
across
a
wide
range
of
excitation
wavelengths,
hence
providing
a
visual
means
to
assess
structural
integrity
as
well
as
qualitatively
verify
protein
removal
[35,61].
Cross-sectional
images
of
the
unloaded
natural
pollen
and
two
SEC
samples
confirm
that
the
processing
steps
effectively
removed
sporoplasmic
contents
from
inside
the
exine
capsules,
whereas
an
extensive
quantity
of
autofluorescent
contents
is
visible
inside
the
natural
pollen
sample.
FITC-labeled
BSA
protein
was
next
loaded
by
vacuum
methods
into
the
samples
as
previously
described,
so
that
the
protein
could
be
visualized
by
CLSM.
Under
equivalent
image
settings,
cross-sectional
slices
were
collected
and
recon-
structed
to
form
a
three-dimensional
representation
of
the
individual
particles.
It
was
observed
that
the
fluorescence
intensity
of
loaded
protein
was
typically
greater
for
SEC
samples
than
for
natural
pine
pollen
grains,
suggesting
that
the
SECs
have
higher
loading
efficiencies
than
natural
pollen.
In
particular,
the
loading
inside
the
central
cavity
appeared
to
be
greater
for
SECs
versus
the
natural
pine
pollen.
Co-localization
of
the
fluorescence
signals
from
the
pollen
exine
wall
(blue
channel)
and
the
loaded
protein
(green
channel)
further
supports
that
the
protein
is
encapsulated
within
the
particles
in
all
three
cases.
To
verify
protein
loading,
absorbance
measurements
were
also
conducted
in
order
to
quantitatively
determine
the
loading
efficiency
of
the
different
samples.
The
vacuum
loading
method
was
utilized,
and
the
loaded
samples
were
extensively
washed
before
measurement.
The
natural
pine
pollen
had
a
loading
efficiency
of
7.9

1.5%,
which
is
comparable
to
other
pollen
species.
This
low
efficiency
is
likely
attributed
to
the
presence
of
sporoplasmic
contents
in
the
central
cavity.
By
contrast,
the
loading
efficiency
of
the
SECs
was
around
three-times
greater,
with
values
in
the
range
of
23–26%
that
demonstrate
excellent
loading
capacity.
The
phosphoric
acid-treated
SECs
had
a
loading
efficiency
of
23.0

2.6%,
whereas
the
hydrochloric
acid-treated
SECs
(with
additional
trypsin
treatment)
had
a
loading
efficiency
of
26.5

3.7%.
These
findings
strongly
support
the
CLSM
results,
and
demonstrate
that
pine
pollen
SECs
can
be
prepared,
which
are
morphologically
intact,
devoid
of
protein
contents,
and
capable
of
efficient
loading.
The
increased
loading
of
the
SECs
can
be
explained
by
the
removal
of
sporoplasmic
contents
(greater
available
loading
volume
per
particle
and
a
higher
number
of
particles
per
unit
mass)
as
well
as
dissolution
of
the
intine
layer,
which
increases
access
to
the
nanoscale
channels
facilitating
protein
encapsulation.
Of
note,
while
loading
appeared
to
be
greater
in
the
air
sacs
for
the
natural
pine
pollen
grains,
the
loading
appeared
to
be
greater
in
the
central
cavity
for
the
SECs.
This
difference
in
loading
properties
indicates
that
chemical
processing
affects
the
molecular
permeability
of
one
or
both
cavity
types.
In
particular,
these
findings
support
that
the
permeability
of
the
central
cavity
in
natural
pine
pollen
grains
is
largely
controlled
by
the
molecular
properties
of
the
intine
layer
and
that
chemical
processing
removes
this
intine
layer
[70].
As
demonstrated
in
this
work,
systematic
investigation
of
chemical
processing
strategies
identified
that
85%
phosphoric
acid
for
5
h
at
70

C
is
optimal
while
other
strategies
are
also
possible.
Conclusion
While
pine
pollen
is
among
the
most
widely
used
pollen
species
in
industrial
settings,
the
development
of
pine
pollen
SECs
for
microencapsulation
applications
has
remained
unexplored.
Here-
in,
we
addressed
this
gap
by
conducting
a
systematic
investigation
aimed
at
identifying
optimal
chemical
processing
strategies
to
extract
pine
pollen
SECs.
Several
processing
parameters
were
tested,
including
acid
type,
processing
duration,
temperature,
and
storage
method,
and
the
results
were
evaluated
by
characterizing
the
morphological
properties
of
resulting
SECs
as
well
as
the
efficiency
of
protein
removal.
Based
on
these
characterization
efforts,
it
was
observed
that
one-pot
acidolysis
with
85%
phosphoric
acid
for
5
h
at
70

C
was
the
optimal
condition
on
account
of
preserving
the
multi-compartment
capsule
architecture
and
successfully
removing
proteins.
It
was
also
possible
to
prepare
SECs
by
utilizing
hydrochloric
acid
together
with
subsequent
enzymatic
treatment,
although
hydrochloric
acid
or
other
strong
acids
alone
were
insufficient
to
effectively
remove
proteins.
The
resulting
SECs
demonstrated
three-times
greater
loading
efficien-
cies
than
the
natural
pollen
grains,
and
CLSM
imaging
demon-
strated
that
the
higher
loading
efficiency
primarily
arises
from
greater
encapsulation
within
the
central
cavity.
This
finding
directly
supports
that
removal
of
sporoplasmic
contents
facilities
higher
loading
efficiencies
of
the
SECs,
as
compared
to
the
unprocessed
pollen
grains.
In
summary,
these
results
outline
a
successful
processing
strategy
to
prepare
SECs
from
multi-
compartmental
pollen
capsules
and
demonstrate
that
the
resulting
capsules
can
be
loaded
with
high
efficiency.
Given
the
wide
range
of
applications
for
pine
pollen
grains,
there
is
excellent
potential
for
applying
these
processing
strategies
to
utilize
pine
pollen
SECs
in
real-life
applications
such
as
drug
delivery
and
taste-masking.
Acknowledgements
This
work
was
supported
by
the
National
Research
Foundation
(NRF-NRFF2011-01),
an
NRF
Competitive
Research
Programme
(NRF-CRP10-2012-07),
and
the
National
Research
Foundation
of
Korea
(KNRF/2-1605-0017).
References
[1]
R.
Sridhar,
R.
Lakshminarayanan,
K.
Madhaiyan,
V.A.
Barathi,
K.H.C.
Lim,
S.
Ramakrishna,
Chem.
Soc.
Rev.
44
(2015)
790.
[2]
M.P.
Adhikari,
R.
Adhikari,
R.G.
Shrestha,
R.
Rajendran,
L.
Adhikari,
P.
Bairi,
R.R.
Pradhananga,
L.K.
Shrestha,
K.
Ariga,
Bull.
Chem.
Soc.
Jpn.
88
(2015)
110 8.
[3]
T.-R.
Su,
Z.-J.
Liao,
M.-C.
Lu,
Y.-J.
Wu,
J.-H.
Su,
Bull.
Chem.
Soc.
Jpn.
88
(2014)
176 .
[4]
M.
Govindarajan,
G.
Benelli,
RSC
Adv.
6
(2016)
59021.
[5]
R.
Scott,
Pollen
Exine—The
Sporopollenin
Enigma
and
The
Physics
of
Pattern,
Cambridge
University
Press,
1994,
pp.
49.
[6]
R.
Wiermann,
S.
Gubatz,
Int.
Rev.
Cytol.
140
(1992)
35.
[7]
R.P.
Wodehouse,
Pollen
Grains.
Their
Structure,
Identification
and
Significance
in
Science
and
Medicine,
Mc
Graw-Hill
Publishing
Co.
Ltd.,
London,
1935.
[8]
P.
Bedinger,
Plant
Cell
4
(1992)
879.
[9]
J.
Brooks,
G.
Shaw,
Grana
17
(1978)
91.
[10]
R.
Wiermann,
F.
Ahlers,
I.
Schmitz-Thom,
Biopolym.
Online
(2001).
[11]
J.
Brooks,
G.
Shaw,
Nature
219
(1968)
532.
[12]
J.
Brooks,
G.
Shaw,
Pollen.
Development
and
Physiology,
Butterworth
&
Co.
Ltd.,
1971.
[13]
E.
Domínguez,
J.A.
Mercado,
M.A.
Quesada,
A.
Heredia,
Sexual
Plant
Reprod.
12
(1999)
171.
[14]
F.
Zetzsche,
O.
Kälin,
Helv.
Chim.
Acta
15
(1932)
412.
[15]
M.S.
Amer,
R.
Tawashi,
Modified
pollen
grains
for
delivering
biologically
active
substances
to
plants
and
animals,
Google
Patents,
1991.
384
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385

[16]
M.S.
Amer,
R.
Tawashi,
Drug
loaded
pollen
grains
with
an
outer
coating
for
pulsed
delivery,
Google
Patents,
1994.
[17]
M.
Couderchet,
J.
Schmalfuß,
P.
Böger,
Pestic.
Biochem.
Physiol.
55
(1996)
189 .
[18]
E.
Domínguez,
J.A.
Mercado,
M.A.
Quesada,
A.
Heredia,
Grana
37
(1998)
93.
[19]
G.
Erdtman,
Svensk
Bot.
Tidskr.
54
(1960)
561.
[20]
K.E.
Espelie,
F.A.
Loewus,
R.J.
Pugmire,
W.R.
Woolfenden,
B.G.
Baldi,
P.H.
Given,
Phytochemistry
28
(1989)
751.
[21]
M.
Hesse,
M.
Waha,
Plant
Syst.
Evol.
163
(1989)
147.
[22]
F.
Zetzsche,
K.
Huggler,
Eur.
J.
Org.
Chem.
461
(1928)
89.
[23]
F.
Zetzsche,
O.
Kälin,
Helv.
Chim.
Acta
14
(1931)
517.
[24]
F.
Zetzsche,
P.
Kalt,
J.
Liechti,
E.
Ziegler,
Journal
für
Praktische
Chemie
148
(1937)
267.
[25]
F.
Zetzsche,
H.
Vicari,
Helv.
Chim.
Acta
14
(1931)
62.
[26]
S.U.
Atwe,
Y.
Ma,
H.S.
Gill,
J.
Control.
Release
194
(2014)
45.
[27]
L.
Wang,
J.A.
Jackman,
W.B.
Ng,
N.J.
Cho,
Adv.
Funct.
Mater.
26
(2016)
8623.
[28]
X.-X.
Wang,
K.
Tian,
H.-Y.
Li,
Z.-X.
Cai,
X.
Guo,
RSC
Adv.
5
(2015)
29428.
[29]
V.N.
Paunov,
G.
Mackenzie,
S.D.
Stoyanov,
J.
Mater.
Chem.
17
(2007)
609.
[30]
S.M.
Alshehri,
H.A.
Al-Lohedan,
A.A.
Chaudhary,
E.
Al-Farraj,
N.
Alhokbany,
Z.
Issa,
S.
Alhousine,
T.
Ahamad,
Eur.
J.
Pharm.
Sci.
88
(2016)
158.
[31]
A.
Diego-Taboada,
S.T.
Beckett,
S.L.
Atkin,
G.
Mackenzie,
Pharmaceutics
6
(2014)
80.
[32]
A.
Diego-Taboada,
L.
Maillet,
J.H.
Banoub,
M.
Lorch,
A.S.
Rigby,
A.N.
Boa,
S.L.
Atkin,
G.
Mackenzie,
J.
Mater.
Chem.
B
1
(2013)
707.
[33]
S.A.
Hamad,
A.F.
Dyab,
S.D.
Stoyanov,
V.N.
Paunov,
J.
Mater.
Chem.
21
(2011)
18018 .
[34]
M.
Lorch,
M.J.
Thomasson,
A.
Diego-Taboada,
S.
Barrier,
S.L.
Atkin,
G.
Mackenzie,
S.J.
Archibald,
Chem.
Commun.
(2009)
6442.
[35]
R.C.
Mundargi,
V.R.
Babu,
V.
Rangaswamy,
P.
Patel,
T.M.
Aminabhavi,
J.
Control.
Release
125
(2008)
193.
[36]
A.
Wakil,
G.
Mackenzie,
A.
Diego-Taboada,
J.G.
Bell,
S.L.
Atkin,
Lipids
45
(2010)
645.
[37]
J.D.
Ackerman,
Abiotic
Pollen
and
Pollination:
Ecological,
Functional,
and
Evolutionary
Perspectives,
Springer,
2000,
pp.
167.
[38]
T.M.
Culley,
S.G.
Weller,
A.K.
Sakai,
Trends
Ecol.
Evol.
17
(2002)
361.
[39]
E.
Duhoux,
Zeitschrift
für
Pflanzenphysiologie
99
(1980)
207.
[40]
S.
Gubatz,
S.
Herminghaus,
B.
Meurer,
D.
Strack,
R.
Wiermann,
Pollen
et
spores
(1986).
[41]
S.
Herminghaus,
S.
Gubatz,
S.
Arendt,
R.
Wiermann,
Zeitschrift
für
Naturforschung
C
43
(1988)
491.
[42]
D.
Southworth,
Am.
J.
Bot.
(1985)
1274.
[43]
J.H.
Park,
J.
Seo,
J.A.
Jackman,
N.-J.
Cho,
Sci.
Rep.
6
(2016)
28017.
[44]
G.-X.
Mao,
L.-D.
Zheng,
Y.-B.
Cao,
Z.-M.
Chen,
Y.-D.
Lv,
Y.-Z.
Wang,
X.-L.
Hu,
G.-F.
Wang,
J.
Yan,
Oxid.
Med.
Cell.
Longev.
2012
(2012).
[45]
B.S.
Hansen,
E.J.
Cushing,
Geol.
Soc.
Ame.
Bull.
84
(1973)
1181.
[46]
B.F.
Jacobs,
Contrib.
Seri.:
Am.
Assoc.
Stratigr.
Palynol.
16
(1985)
155.
[47]
R.N.
Mack,
Pollen
Size
Variation
in
Some
Western
North
American
Pines
as
Related
to
Fossil
Pollen
Identification,
Washington
State
University
Press,1971.
[48]
W.S.
Ting,
196 6.
http://agris.fao.org/agris-search/search.do?recordID=US201
300317259.
[49]
D.
Attenborough,
M.
Salisbury,
N.
Nightingale,
I.
Haynes,
M.
Elsbury,
J.
Payne,
S.
Olive,
The
Private
Life
of
Plants,
JSTOR,
1995.
[50]
G.
Bohne,
H.
Woehlecke,
R.
Ehwald,
Ann.
Bot.
96
(2005)
201.
[51]
G.
Bohne,
E.
Richter,
H.
Woehlecke,
R.
Ehwald,
Ann.
Bot.
92
(2003)
289.
[52]
S.
Barrier,
University
of
Hull,
2008.
http://ethos.bl.uk/OrderDetails.do?uin=uk.
bl.ethos.564124
(thesis).
[53]
S.
Barrier,
A.
Diego-Taboada,
M.J.
Thomasson,
L.
Madden,
J.C.
Pointon,
J.D.
Wadhawan,
S.T.
Beckett,
S.L.
Atkin,
G.
Mackenzie,
J.
Mater.
Chem.
21
(2011)
975.
[54]
K.S.
Osthoff,
R.
Wiermann,
J.
Plant
Physiol.
131
(1987)
5.
[55]
A.
Armentia,
A.
Quintero,
A.
Fernandez-Garcia,
J.
Salvador,
J.
Martin-Santos,
Ann.
Allergy
64
(1990)
49.
[56]
G.
Freeman,
Ann.
Allergy
70
(1993)
491.
[57]
S.
Varis,
J.
Reiniharju,
A.
Santanen,
H.
Ranta,
P.
Pulkkinen,
Grana
50
(2011)
129.
[58]
M.E.H.
Bashir,
J.H.
Lui,
R.
Palnivelu,
R.M.
Naclerio,
D.
Preuss,
PLoS
One
8
(2013)
e57566.
[59]
A.J.
Boasman,
The
University
of
Hull,
2003.
http://ethos.bl.uk/OrderDetails.do?
uin=uk.bl.ethos.507478
(thesis).
[60]
R.C.
Mundargi,
M.G.
Potroz,
S.
Park,
H.
Shirahama,
J.H.
Lee,
J.
Seo,
N.J.
Cho,
Small
(2015).
[61]
R.C.
Mundargi,
M.G.
Potroz,
J.H.
Park,
J.
Seo,
E.-L.
Tan,
J.H.
Lee,
N.-J.
Cho,
Sci.
Rep.
6
(2016).
[62]
M.G.
Potroz,
R.C.
Mundargi,
J.H.
Park,
E.-L.
Tan,
N.-j.
Cho,
J.
Visualized
Exp.
(2016)
e54768.
[63]
A.
Diego-Taboada,
P.
Cousson,
E.
Raynaud,
Y.
Huang,
M.
Lorch,
B.P.
Binks,
Y.
Queneau,
A.N.
Boa,
S.L.
Atkin,
S.T.
Beckett,
J.
Mater.
Chem.
22
(2012)
9767.
[64]
R.C.
Mundargi,
M.G.
Potroz,
J.H.
Park,
J.
Seo,
J.H.
Lee,
N.-J.
Cho,
RSC
Adv.
6
(2016)
16533.
[65]
S.
Knoche,
J.
Kierfeld,
Phys.
Rev.
E
84
(2011)
046608.
[66]
D.
Sen,
J.
Bahadur,
S.
Mazumder,
G.
Verma,
P.
Hassan,
S.
Bhattacharya,
K.
Vijai,
P.
Doshi,
Soft
Matter
8
(2012)
1955.
[67]
R.M.
Parker,
J.
Zhang,
Y.
Zheng,
R.J.
Coulston,
C.A.
Smith,
A.R.
Salmon,
Z.
Yu,
O.A.
Scherman,
C.
Abell,
Adv.
Funct.
Mater.
25
(2015)
4091.
[68]
S.A.
Nabavi,
G.T.
Vladisavljevic,
S.
Gu,
V.
Manovic,
Langmuir
32
(2016)
9826.
[69]
F.
Higgins,
G.
Ho,
Agric.
Wastes
4
(1982)
97.
[70]
N.
Wenda,
H.
Woehlecke,
T.
Detloff,
D.
Lerche,
Chem.
Ing.
Tech.
84
(2012)
309.
A.K.
Prabhakar
et
al.
/
Journal
of
Industrial
and
Engineering
Chemistry
53
(2017)
375–385
385