Nanoparticle suspensions enclosed in methylcellulose: a new approach for quantifying nanoparticles in transmission electron microscopy

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Nanoparticles are of increasing importance in biomedicine but quantification is problematic because current methods depend on indirect measurements at low resolution. Here we describe a new high-resolution method for measuring and quantifying nanoparticles in suspension. It involves premixing nanoparticles in a hydrophilic support medium (methylcellulose) before introducing heavy metal stains for visualization in small air-dried droplets by transmission electron microscopy (TEM). The use of methylcellulose avoids artifacts of conventional negative stain-TEM by (1) restricting interactions between the nanoparticles, (2) inhibiting binding to the specimen support films and (3) reducing compression after drying. Methylcellulose embedment provides effective electron imaging of liposomes, nanodiscs and viruses as well as comprehensive visualization of nanoparticle populations in droplets of known size. These qualities facilitate unbiased sampling, rapid size measurement and estimation of nanoparticle numbers by means of ratio counting using a colloidal gold calibrant. Specimen preparation and quantification take minutes and require a few microliters of sample using only basic laboratory equipment and a standard TEM.

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received: 24 December 2015

accepted: 14 April 2016

Published: 04 May 2016

# Nanoparticle suspensions enclosed in methylcellulose: a new approach for quantifying nanoparticles in transmission electron microscopy

Christian Hacker 1,† , Jalal Asadi 1 , Christos Pliotas 2 , Sophie Ferguson 1 , Lee Sherry 3 ,
Phedra Marius 2 , Javier Tello 1 , David Jackson 3 , James Naismith 2 & John Milton Lucocq 1

Nanoparticles are of increasing importance in biomedicine but quantification is problematic because
current methods depend on indirect measurements at low resolution. Here we describe a new highresolution method for measuring and quantifying nanoparticles in suspension. It involves premixing
nanoparticles in a hydrophilic support medium (methylcellulose) before introducing heavy metal
stains for visualization in small air-dried droplets by transmission electron microscopy (TEM). The use
of methylcellulose avoids artifacts of conventional negative stain-TEM by (1) restricting interactions
between the nanoparticles, (2) inhibiting binding to the specimen support films and (3) reducing
compression after drying. Methylcellulose embedment provides effective electron imaging of
liposomes, nanodiscs and viruses as well as comprehensive visualization of nanoparticle populations
in droplets of known size. These qualities facilitate unbiased sampling, rapid size measurement
and estimation of nanoparticle numbers by means of ratio counting using a colloidal gold calibrant.
Specimen preparation and quantification take minutes and require a few microliters of sample using
only basic laboratory equipment and a standard TEM.

Nanoparticles are of growing importance in biology, medicine, pharmacology, cosmetics and environmental
studies. Indeed a range of synthetic nanoparticles are used increasingly as drug carriers 1,2, diagnostic and analytic
tools 3, therapeutic reagents 4–7, or as platforms for membrane proteins in structural and functional studies 8,9 . The
materials used for synthesis can be in “condensed” form, as in metallic colloids or carbon nanotubes or composed
of “soft” biological substances such as the lipids used to make liposomes, nanodiscs or bicelles. Nanoparticles
can also occur naturally with those of particular significance being pathogenic viruses, important for diagnosis
or vaccine development, or the extracellular vesicles called exosomes that are emerging as markers of human
disease 10 . In many of these applications quantitative monitoring of nanoparticles is essential and demands rapid,
accurate and sensitive quantification on the smallest possible samples at high-resolution. The primary morphological parameters of interest are particle number (concentration), particle size and the degree of aggregation 11 .

Currently nanoparticle quantification is based mainly on biophysical approaches 12 . These include
nanoparticle-tracking analysis (NTA) 12–14, flow cytometry 15, dynamic light scattering (DLS or photon correlation microscopy) 16,17, small angle X ray analysis and resistive pulse sensing. However these methods are largely
indirect in nature and can be prone to systematic errors, especially when nanoparticles vary in size or aggregation state. A more direct approach for characterizing nanoparticles is electron microscopy 11, which provides
nanometer-scale visualization of particles and their morphology. Current EM technologies suited to nanoparticle
investigation include TEM, focused ion beam scanning EM (FIBSEM) and scanning EM (SEM) 11 . In particular
section based methods such as TEM and FIBSEM are especially powerful when applied to nanoparticles that
are dispersed in solid matrices where estimating particle numbers and matrix volume are relatively straightforward and the aggregation state is well preserved. However a significant problem occurs when characterizing

1School of Medicine, University of St Andrews, St Andrews, Fife, UK. 2Centre for Biomolecular Sciences, University
of St Andrews, St. Andrews, Scotland. 3 Biomedical Sciences Research Complex, University of St Andrews, North
Haugh, St Andrews, United Kingdom. † Present address: Bioimaging Centre Geoffrey Pope building College of Life
and Environmental Sciences University of Exeter Stocker Road EX4 4QD Exeter, UK. Correspondence and requests
[for materials should be addressed to J.M.L. (email: jml7@st-andrews.ac.uk)](mailto:jml7@st-andrews.ac.uk)

nanoparticles in liquid phase for which there are a number of possible solutions. The first is to visualize the
dispersion directly in the liquid state, involving encapsulation in specialized containers that can be inserted into
the vacuum needed for electron microscopes to function 11 . However, this technology is still at an early stage
and there are concerns about effects of the electron beam on the nanoparticles. Another solution is to capture or
solidify the nanoparticle suspension by rapid freezing and visualize in the frozen state – a technique referred to as
cryo-EM. Advantages of this approach include preservation of aggregation state and hydration 18, as well as a lack
of a requirement for heavy metal contrast. Therefore cryo-EM represents an optimum approach for qualitative
observation 19 although quantification remains technically challenging because variations in specimen quality
make proper exhaustive analysis or proper sampling difficult. A third solution for visualizing nanoparticles that
are dispersed in liquid phase is to dry the particle suspension in such a way as to avoid morphological or chemical
modification through denaturation, and to avoid artifactual aggregation.

A classical technique for structural analysis that uses drying of nanoparticle suspensions is negative stain
TEM 20 . Here nanoparticles are absorbed onto an EM support and heavy metal stain applied. This method is
the mainstay of high resolution analysis revealing structural details of soft nanoparticles (lipid membranes and
proteins) as well as nanoparticles composed of condensed, “hard”, materials such as metallic particles, carbon or
plastics. Importantly negative stain has not come into general use for quantifying nanoparticles, because it can
generate artifacts such as particle collapse, aggregation, and non-uniform deposition 20 . A key problem is that the
adsorption of nanoparticles to the EM support film cannot be easily controlled, resulting in a poor representation
of the whole particle population for sampling. Decades ago, attempts to overcome this problem used calibration
beads that were applied to EM grids by centrifugation, spraying or drying along with the nanoparticles 21–26 ; but
the results had low precision, were lengthy and relied on quantitative assumptions about the bead preparations.
In the interim TEM has been viewed as slow and technically demanding, producing results of variable quality 15 .

With these problems in mind we developed a simple TEM method that uses embedment of aqueous suspensions of nanoparticles within hydrophilic films composed of methylcellulose. Distinct from the negative stain
approach, the method disperses nanoparticles in standard volumes of methylcellulose containing uranyl acetate
stain before drying. The method restricts aggregation and patchy distributions, and hinders particle collapse
while retaining fine structure. Comprehensive visualization of the nanoparticle population allows unbiased sampling for measurement and characterization of a range of nanoparticle types. Methylcellulose also allows stabilization of metal calibration particles in the hydrophilic films, facilitating estimation of nanoparticle numbers/
concentration with precision and sensitivity. Preparation and quantification protocols are achieved in minutes
and require only standard laboratory and imaging equipment.

Results and Discussion
Methylcellulose films. Previously methylcellulose films containing electron dense stains have been used
extensively for embedding ultrathin cryosections 27–29 and also for visualizing isolated vesicular organelles but
only afer t their adsorption to EM supports 30 . Our aim was to test whether methylcellulose could stabilize suspensions of nanoparticles, so that standardized volumes of hydrophilic films could be mounted for studying and sampling the whole nanoparticle population in the TEM. Nanoparticle suspensions were first mixed with a solution
of methylcellulose before adding a heavy metal contrasting agent (uranyl acetate). The mixture was then applied
to the center of a support plastic-coated TEM grid as a 0.5 μ l droplet containing 0.15% methylcellulose and 0.03%
uranyl acetate (Fig. 1a). Once dried, the film formed a circular spot, which was thicker near the rim. Such a “coffee
ring” distribution is a common effect observed during drying of fluid droplets 31,32 . Importantly we found that
structural display of liposomes, viruses and nanodiscs (Fig. 1b–i) was consistent across the whole droplet even in
the thicker peripheral parts of the droplet and most nanoparticle preparations tested remained aggregate free. We
found larger droplets (i.e. > 0.5 μ l) dried to form regions of high electron density, obscuring some particles, while
smaller volumes produced films with patchy contrast and marked “collapse” of vesicular structures. Visualizing
all nanoparticles in the droplet was an important prerequisite for unbiased sampling, accurate measurement and
enumeration of the particles in the droplet (described later).

Interestingly, the uranyl acetate concentration used was much more dilute than that commonly used for cryosection contrasting in methylcellulose 27–29 . The staining mechanism likely involves local adsorption of the heavy
metal onto nanoparticles rather than conventional “negative” staining in which biological structures are highlighted by surrounding lakes of stain. This “positive” staining of nanoparticles could provide novel structural
information, especially if combined with tomographic analysis of particle structure.

Nanoparticle suspensions dried in droplets of methylcellulose/uranyl acetate appeared largely free of aggregates, which indicated that methylcellulose prevented interactions by adhering to particle surfaces as an inert
coating 33 . This was further tested using colloidal gold particles, which are well known to aggregate in the presence
of cations. When colloidal gold nanoparticles were added to 1% methylcellulose and phosphate buffered saline
added, the gold particles failed to aggregate substantially (as indicated by the lack of a color change from red to
blue/black). In addition this screening effect appeared to reduce binding of nanoparticles to the plastic EM grid
support film as evidenced by the strong inhibition of liposome binding to the grid support (Fig. 2a).

The possible mechanisms by which MC prevents particle-particle and particle surface interactions in solution
and during drying are of interest. There is a large literature showing that polymers bind to metallic or plastic surfaces via short-range forces (van der Waals), with longer polymers binding tighter than shorter ones (reviewed
in ref. 34). In the standard model of particle-particle interactions, particle surface charges restrict aggregation by
electrostatic repulsion, but on addition of electrolytes the surface charges can be neutralized resulting in aggregation. Importantly when hydrophilic amphiphilic polymers are bound to the particles, the resulting screening
effect allows the nanoparticles to take on properties of the polymer and thereby decreasing the impact of particle
attraction and reducing salt-induced aggregation 34 . Methylcellulose is a neutral water-soluble cellulose derivative
produced by heterogeneous methylation of semi-crystalline cellulose. Previous work demonstrating adsorption

Figure 1. Nanoparticles in methylcellulose films imaged in TEM. ( a ) The stain mix contained 1.5 μ l of 1%
methylcellulose, 7.5 μ l of nanoparticle suspension and 1 μ l of 0.3% uranyl acetate (uranyl acetate was added after
dialysis if required; see text and Fig. 3). 0.5 μ l was loaded onto a standard pioloform coated EM grid support
and allowed to dry before examination in the TEM. ( b - i ) A range of nanoparticles imaged at low magnification
( b - e ) and high magnification ( f - i ). Scale bars ( b - e ) 100 nm and ( f - i ) 50 nm.

of methylcellulose to silica particles surfaces supports this view 33 . It is therefore likely that MC coats the surfaces
involved in TEM processing, preventing particle aggregation as well as adsorption/distortion at the grid support
surface. It appears that MC acts in this way throughout the specimen preparation and drying processes, until the
MC becomes solid.

In conventional negative stain, particles adhere to the grid support and assume preferred orientations after
drying. We argued that nanoparticles embedded in the methylcellulose droplet as a suspension should now
demonstrate freedom to orientate during drying. To test this we prepared nanodiscs that were 12 nm in diameter 35
and embedded them in methylcellulose films of varying final thickness before assessing their orientations.
Nanodiscs are synthetic lipoprotein nanoparticles used as carriers for membrane proteins. In the thinnest films
(made with 0.1% methylcellulose) most of the nanodiscs were oriented “enface” to support film after drying
(Fig. 2b). However in thicker films (made with 0.3–0.5% methylcellulose), a range of tilted orientations became
apparent, showing nanodiscs were increasingly free to orientate during the drying process with increasing film
thickness. To further test whether other asymmetric nanoparticles assume multiple orientations in MC droplets, we embedded gold nanorods in films of varying thickness and measured their projected profile lengths in
TEM images (Fig. 2c). In the thinnest central regions of droplets, made with 0.1% MC, nanorod profiles were on
average longer (mean 46.59 nm coefficient of error 2.5%, n = 45) than those measured in the thicker rim in 0.1%
MC droplets (mean 39.87 nm coefficient of error 3.1%, n = 52) or the central region of film made with 0.5% MC
(mean 36.6 nm coefficient of error 3.2%, n = 38). Tilting analysis in TEM confirmed that shorter rod profiles
found in thicker films were at orientations tilted relative to the horizontal (not shown); by comparison nanorods
in thinner parts of the film lay uniformly horizontal to the MC film (statistical comparison of nanorod lengths
are shown in the legend to Fig. 2c). We also obtained evidence that nanoparticle suspensions are distributed

Figure 2. Methylcellulose restricts binding to the substratum, allows freedom of orientation and
prevents substantial collapse of nanoparticles. ( a ) Binding to substratum. Liposomes were incubated in
1% methylcellulose or water before adsorbing the mix to an EM grid, contrasting, and density of liposomes
determined. ( b ) Orientation. 12 nm diameter nanodiscs were mixed with methylcellulose solution varying
between 0.1% or 0.5% before adding uranyl acetate. 0.5 μ l droplets were loaded onto EM grids and dried before
classifying orientation by TEM (flat, tilted and upright). Nanodiscs become progressively tilted with increasing
concentration of methylcellulose. Bars, 7 nm. ( c ) Gold nanorods were mixed with 0.1% or 0.5% MC. 0.5 μ l drops
were loaded onto EM grids and nanorods situated in either the rim or center were measured. Data are presented
as cumulative fractions. Nanorods in the thinnest films (0.1% MC center) were longer compared to those in
thicker films at the 0.1% MC rim (Kolmogorov-Smirnov test, P 0.001; n 45 and 52 data points respectively)
or the 0.5% MC center (Kolmogorov-Smirnov test, P 0.01; n 45 and 38 data points respectively). The data are
consistent with increased freedom to rotate in thicker MC films. ( d, e ) Tilt analysis of vesicle compression in
methylcellulose films. Vesicles containing (MscL) pore protein were embedded in methyl cellulose as described
in the text and the degree of flattening (oblate spheroid; model) estimated from minor and major axes measured
after 50° specimen tilt. Data are from the whole population (All), n = 38, 20–30 nm vesicles, n = 16 and
50–70 nm vesicles, n = 10. Bar = 50 nm.

Figure 3. Microdialysis procedure using semipermeable cellulose kidney dialysis capillaries. ( a ) Three to
five μ l of sample was drawn into the kidney dialysis straw by capillary action. ( b ) Ends of the straw were sealed
using an assembly made from interlocking truncated micropipette tips. ( c ) The assembly was transferred into
the dialysate. ( d ) The dialyzed sample was extruded onto parafilm and added to methylcellulose/uranyl acetate
mix before loading and drying on an EM support grid.

throughout the thickness of dried MC droplets. After drying droplets containing mixes of gold particle calibrant
and MC, gold particles in the droplet rim appeared to form clusters/aggregates. However these clusters could be
clearly separated by tilting the EM grid relative to the electron beam demonstrating that the nanoparticles spread
through the thickness of the droplet film in the vertical direction.

Published data predicts significant collapse of negatively stained synaptic vesicles. For example the data of one
study indicates a height to width ratio of 0.25 for 47 nm vesicles 36 . We next tested for collapse of nanoparticles
suspended in methylcellulose using liposomes that had spherical morphology. These liposomes carried the protein channel MscL 35, and as previously shown for the mechanosensitive ion channel MscS 35,37–39, it allows for the
import of water into the vesicles making them appear turgid and pseudo-spherical. The degree of flattening after
drying was evaluated by comparing images at 0 and 50° tilt, with consequent modeling of vesicles as oblate spheroids. MscL vesicles had an aspect ratio (height to width) of 0.66 after drying. Overall at this level of compression
the volume equivalent vesicle sphere had a diameter, which was 0.88 of the spheroid major axis measurement
indicating an overestimation of vesicle size of approximately 12% using the methylcellulose method. Interestingly
larger turgid vesicles appeared more flattened (50–70 nm diameter (26% of the population); aspect ratio 0.56)
than smaller ones (20–30 nm (34% of the population); aspect ratio 0.8; Fig. 2c). We conclude that methylcellulose
suspensions support vesicles better than negative stain. Another interesting approach to reduce collapse of structures could be cryo-EM, but this technique was not investigated further because, as with negative stain, visualization of particle populations is patchy making valid sampling and quantitation problematic 40 .

De-salting nanoparticle preparations using a novel microdialysis system. Biological nanoparticle
preparations contain salts that interfere with electron stains and/or may crystalize during drying, thereby obscuring nanoparticles. The applicability of the method was further enhanced with a microdialysis system (Fig. 3) that
makes use of cellulose dialysis straws obtained from kidney dialysis units (internal diameter of the capillaries was
approximately 200 microns with a molecular weight cut-off of 10,000 Da). A mix of methylcellulose solution and
colloidal gold calibrant was combined with nanoparticle solution and three to five microliters loaded into straws
by capillary action before sealing (Fig. 3a). After dialysis against deionized water, the dialysate was extruded and
heavy metal stain added before loading the droplet onto an EM grid (Fig. 3b–d). After dialysis, neither aggregation of gold particles or NPs, nor precipitation of stain was observed. Importantly we did not observe blebbing of
liposomes or virus under the low molarity conditions. This novel procedure allowed microliter samples containing nanoparticles to be processed for quantitation in a variety of culture supernatants, laboratory samples and
body fluids.

Quantifying nanoparticle size. We next developed a method for particle sizing by combining the resolution of TEM with a sampling based stereological method. To increase throughput the method was applied to a live
digital camera display during x/y scanning across the grid. The principle was tested using liposomes that are used
widely for drug/reagent delivery and as carriers for study of protein function 41 . Liposomes embedded in methylcellulose droplets were selected during x/y scanning by applying unbiased counting rules to a scanning-band
(Fig. 4) 42,43 . The selected liposomes were then translocated through a systematically spaced array of dots that was
placed across the digital camera display (Fig. 4a; see ref. 42). In this system the number of intercepts between each
translocating liposome with the upper dot edges can be used to estimate particle caliper diameter unbiasedly
(where Cd (caliper diameter) = Ilip (number of intercepts with the liposome) × d (spacing of the dot edges)); given

Figure 4. Liposome sampling and size estimation. Liposomes were sampled using a scanning band to which
an unbiased counting rule was applied ( a ). All particles passing completely between the lines or intersecting the
acceptance line (green) were selected for analysis. Particles intersecting the forbidden line (red) were excluded.
Selected particles were translocated through size a systematic series of dots the upper edge of each tracing a
scanning line. The particle caliper diameter was calculated from the number of intersection of dot edges with
the nanoparticle and the real spacing of the dots as described in the text. Data are from three experimental runs
from the same liposome preparation ( b ). Approximately 100 vesicles were counted per experiment; error bars
are standard error of the mean (n = 3; each estimation took approximately 10 minutes).

random positioning of the particles and/or line array. For each preparation the results from approximately 100
liposomes were grouped into at least 10, 30 nm size bands (Fig. 4b). Evaluation of each liposome preparation took
10 minutes and over three experiments the average coefficient of error for size estimates over all size bands under
210 nm diameter was 8%. The distributions from the three experiments were not distinguishable as determined
by contingency/Chi square analysis (p > 0.25, df 16).

Quantifying gold calibrant. The principle was to mix gold particle calibrant (15 nm gold particles at known
concentration and prepared with the citrate reduction method 34,42 ) with nanoparticles of interest and count the
ratio of gold to nanoparticles. As already noted, salt-induced aggregation of colloidal gold 34 was restricted in 1%
methylcellulose, which allowed the calibration solution to be premixed with nanoparticle suspensions in laboratory buffers and body fluids before removal of the salts and addition of heavy metal stain.

Known volumes of calibrant were included in methylcellulose suspension and 0.5 μ l of this mixture loaded
onto an EM grid. The dried circular droplets were then sampled using a systematic uniform random (SUR)
pattern 45,46 of imaging windows across the whole droplet. The use of SUR sampling 45,46 ensures unbiasedness and
optimal efficiency even with increased particle density in the droplet rim. Indeed, SUR provides better precision
and is more rapid to implement than simple random sampling. The total number of gold particles in 0.5 μ l was
computed from the number of particles counted in the windows multiplied by the ratio of the droplet area to the
collective sampled area in the windows. The concentration of gold particles in the calibration preparation used
for quantifying the nanoparticles used in this study (liposomes and influenza virus) was found to be 4.4 × 10 10
particles ml −1 (coefficient of error, 6.47%; n = 8; see Supplementary Figure 1).

Figure 5. Quantification of nanoparticles using gold particle calibration. ( a ) During scanning gold
particles and test nanoparticles were imaged “live” using a digital camera and counted using a scanning band
as described in the text and in Fig. 4. The tip of the green arrow traces the acceptance line (dashed green) and
red arrow the forbidden line (dashed red) during translocation of the specimen. The liposome (nanoparticle)
concentration was estimated from the ratio of golds to liposome (nanoparticle) multiplied by the gold
calibrant concentration. ( b ) Estimates of gold/nanoparticle ratio from single equatorial scans closely match
those obtained from multiple SUR scans. Number estimates were obtained from the same grid but in different
locations for each type of sample (n = 3, mean value and bars standard error of mean). Scale bar, 500 nm.

To further validate the counting method we compared estimates of concentration in gold calibrants with an
independent measure of gold particle number based on 3D models. A popular model based approach for estimating gold particle concentration assumes spherical shape of particles. However, this underestimates gold particle
number because gold colloidal particles are a mix of crystalline structures such as the truncated octahedrons that
are the favored for this particle size 47 . Indeed the crystalline morphology of our gold calibrant was supported
by the observation of particles with multifaceted profiles and densities that were interpreted as due to electron
scattering by crystal vertices. By measuring the maximal diameters of gold particles (circumscribed sphere diameters), we could then apply known ratios of sphere/crystal volumes for truncated octahedrons. We estimated a
particle concentration of 1.61 × 10 12 per ml. By comparison the droplet counting method from the same gold colloid produced a mean value of 1.688 × 10 12 per ml (coefficient of error, 2.93%; n = 3; see supplementary materials
for details). It is important to note that there are few other methods for reliable counting of small nanoparticles.
We attempted to count the gold particles or liposomes using flow cytometry and nanoparticle-tracking analysis
but were unable to produce consistent results.

Quantifying nanoparticles–gold calibration/nanoparticle ratio. The use of the gold particle calibrant for counting liposomes by ratio estimation is illustrated in Fig. 5. Liposomes were mixed with methylcellulose/gold calibration solution and then uranyl acetate added (dialysis was not necessary here because
the liposome carrier was compatible with heavy metal staining). 0.5 μ l was loaded onto support grids before
air-drying and neither liposomes nor gold particles showed evidence of aggregation. Also they did not interact

Figure 6. Quantification of liposomes. Scaling between liposome dilution and estimates of liposome particle
number. In each case, approximately 100 each of gold calibration and liposomes were counted (time for each
scan 10–15 minutes). Values are means of 4 separate dilution series derived from the same liposome preparation
and error bars are coefficients of variance.

substantially with each other (see Fig. 5a; < 1% of gold particles were located at liposomes) and did not appear to
segregate from each other in different areas of the droplet.

Gold and liposomes (nanoparticles) were counted on a live digital camera display to increase throughput
(Fig. 5a). The particles were sampled by applying unbiased counting rules 45,46 to a scanning band (Figs 4 and 5a)
and all sampled gold and liposomes particles “exiting” the band were counted (Fig. 5b). Initially approximately
10 scans were positioned SUR across the whole droplet (SUR sampling is by definition unbiased). As already
mentioned SUR is more efficient than random sampling especially when distributions are heterogeneous as is
expected in the coffee ring effect 31,32 . However to improve efficiency further, we tested whether a single scan
placed across the grid equator, would provide results that closely match the results obtained using this SUR
approach; again counting approximately 100 or more gold or nanoparticles (i.e. liposomes; Fig. 5b). A single scan
was much quicker to carry out and took approximately 10 minutes providing a mean estimate that was within
1.2% of the SUR value. Initially when comparing SUR scans with a single equatorial scan we counted an average of
364 gold particles and 60 liposomes per experiment. However it is a general principle in stereology that 100–200
events are usually sufficient for workable precision and if one particle type is more abundant than the other the
counts can be balanced by reducing the scanning bandwidth for the most abundant particle type (the overall
quantity of counts can also be altered by adjusting the magnification). One possibility is that the different particle
types are distributed differently during drying especially in the droplet periphery 48,49 . Theoretically this effect
could be detected by differences in the estimates using SUR and equatorial sampling because equatorial sampling
favors the more central populations while SUR does not. Importantly the strong similarities in results from these
different sampling strategies indicate that separation effects are minor or absent. It is possible that methylcellulose
could reduce such separation effects by reducing the surface tension 49 .

We next tested precision of number estimates across a range of dilutions. The dilutions covered approximately
4 orders of magnitude and the average coefficient of variation was 6.3%, although at the lowest concentrations
the precision ranged from 1.9 to 4.1% (Fig. 6). Scaling was close to linear (Correlation coefficient R 2 = 1). When
the liposome concentration was above 10 7 /ml counting took approximately 10–15 minutes. To test the limits of
sensitivity we scanned the droplet in the rim only where the film was thickest and presented the highest concentration of nanoparticles. On this basis we found an estimable concentration of nanoparticles to be approximately
5 × 10 6 ml −1 .

Quantifying a test sample–influenza A virus. To test the feasibility of the counting method we estimated nanoparticle numbers in three independent preparations of influenza A virus particles. These preparations contained phosphate buffered saline and were mixed with first methylcellulose/gold calibrant before dialysis
against deionized water. Uranyl acetate was then added before droplet loading and drying. Ratio counting of
approximately 100 gold and 100 virus particles was carried out using single equatorial scans (Fig. 7; each count
required between 5 and 10 minutes). Infectivity was also assayed in parallel using plaque forming unit (PFU)
assays, allowing the particle to infectivity ratio to be determined. The particle/infectivity ratio was estimated to be
69.9:1 (coefficient of variation for particle number was 15.4% and for PFU was 47.5%; demonstrating enhanced
precision of methyl cellulose embedment over the PFU method). Methylcellulose embedment proved to be much
more precise and rapid than the widely used haemagglutination (HA) assay 50 which is indirect, well known to
be imprecise and relies on assumptions about thresholds of particle aggregation 51 . Our estimates of virus particle
number indicate the ratio of particles to infective units might be substantially higher than indicated by previous
estimates 52 .

Figure 7. Influenza A virus quantification. ( a ) The virus preparation was mixed with gold/methylcellulose
calibration solution, dialyzed by the microdialysis method (see Fig. 3) and embedded in methylcellulose/uranyl
acetate. Estimates of virus particle number using the suspensions in methylcellulose method were compared
with plaque forming units (PFU assay), determined on the same samples (n = 3; error bars, standard error
of the mean). ( b ) TEM image showing lack of gold (arrows) or virus particle interaction/aggregation. Scale
bar = 100 nm.

Figure 8. Overview of suspension in methylcellulose method. Nanoparticle (NP) suspension is added to
gold calibrant (Au)/methylcellulose (MC) to make a calibration mix (NP/Cal mix). If necessary, NP/Cal mix is
dialyzed in microdialysis straws and extruded before adding heavy metal stain (uranyl acetate). 0.5 μ l is loaded
onto the plastic coated TEM support grid for counting, sizing and further characterization.

Workflow summary. Figure 8 summarizes the workflow for quantifying an unknown nanoparticle preparation. First the preparation is mixed 1:1 with gold/methylcellulose calibration. If necessary microdialysis of
3–5 μ l of the nanoparticle/calibration mix against uranyl acetate-compatible solutions can be performed in cellulose straws. Uranyl acetate is then added to a final concentration of 0.03%, and 0.5 μ l is then loaded onto a
pioloform-coated EM grid before air-drying. At the TEM, a single equatorial scan across the center of the droplet
is used to size nanoparticles or estimate the ratio of calibrant particles/nanoparticle according to Figs 4 and 5. The
width of particle sampling bands and, if required magnification, are adjusted according to the abundance of nanoparticles/calibration gold to count 100–200 of each (the counts in the smaller band are multiplied by the ratio of
the large band size to the small band size, to allow comparison).

Conclusions
In conclusion we have developed a rapid method for counting and sizing nanoparticles at high resolution,
improving on widely used biophysical methods. We consider the high precision and efficiency to be built on
two main advantages. The first is the property of methylcellulose to maintain nanoparticles in homogeneous
suspension, reducing aggregation and artifacts while presenting the whole population for quantitative analysis.
The second is the use of rigorous and efficient sampling tools for quantitation that reduce the workload. In future
further improvements in efficiency could be made by using automated feature recognition to remove reliance on
operator-based counting/sizing. We recently developed algorithms for identifying and sizing particles automatically 53 and their use in combination with the present method would improve throughput. Another improvement
would be the labeling of molecular nanoparticle components, which is a powerful method for categorizing nanoparticles and membrane bound vesicles 54 . We have not extended the current technique in this way but affinity
labeling techniques using multiple sizes or shapes of electron dense markers 55 would provide correlation of size
or detailed structure with molecular composition. Compared to particle tracking analysis, the method improves
sensitivity by two orders of magnitude and reduces sample size from 100 s to just a few microliters. By incorporating microdialysis, microliter samples from culture supernatants or body fluids can be analyzed. Preparation,
counts or size distributions take a few minutes and require little training, and use standard electron microscopy to
provide nanometer resolution of structural features. Although our results show that the current method is useful
for counting and sizing of nanoparticles they do not at present detect the aggregation state of the particles before
they were mixed with MC. Given that aggregation state is a key parameter in nanoparticle research it will be of
interest to assess the effects by independent measures of particles aggregation such as cryo-EM or DLS 18 .

The method appears to be suitable to a wide range of nanoparticle types from those composed of condensed
materials with inherent electron contrast (metallic colloids or plastic beads), to those composed of soft biological
materials for which electron dense staining may then be a prerequisite. The method was tested for use on nanoparticles dispersed in aqueous media, which were made compatible with the use of electron stains by prior dialysis. Future studies will reveal whether other embedding media or solvents can be used, opening up the droplet
method to a wider range of nanoparticle types.

Embeddment in MC droplets should now facilitate direct analysis of tiny samples of synthetic and naturally
occurring nanoparticles widely used in laboratory studies, cosmetics, pharmacology, theranostics and nanotoxicology. Specifically these applications could include: (1) counting and sizing of commercial metal colloids and
plastic/polymer beads in quality control and monitoring during synthesis and use, (2) the characterization of
liposomes used in drug delivery and cosmetics, (3) preparation of multifunctional nanoparticle preparations in
therapeutics, (4) diagnosis of viruses and preparation of viruses in vaccine development as well as (5) the study
of extracellular vesicles such as exosomes in laboratory samples and body fluids, thereby facilitating their use as
markers in human disease.

Methods
Cells, Virus and Plaque Assays. MDCK and 293T cells were maintained in Dulbecco’s modified Eagle’s
medium (DMEM, supplemented with 10% fetal calf serum; Invitrogen, Paisley, UK) at 37 °C in a 5% CO2 atmosphere. Influenza A virus/WSN/33 wild-type (rWSN wt) was generated using plasmid-based reverse genetics as
previously described 56 . Briefly, 293T cells were transfected with eight virus genome-encoding plasmids (pHH-21based; encoding genes under the control of the human RNA polymerase I promoter) and four protein expression
plasmids encoding the genes for the viral polymerase subunits (PB1, PB2, PA) as well as nucleoprotein (nanoparticle) under control of a CMV polymerase II promoter. At 16 h post-transfection, cells were co-cultured with
MDCK cells in serum-free DMEM containing 2.5 μ g/mL N-acetyl trypsin (Sigma Aldrich, Dorset, UK). Viruscontaining supernatant was harvested three days post-transfection, viruses propagated twice through MDCK
cells, followed by plaque assay titration on MDCK cells. MDCK cells in six-well plates were infected with serial
10-fold dilutions of each virus in serum-free DMEM for 1 h at 37 °C. Cells were overlaid with DMEM–1% agarose
supplemented with 2 μ g/ml N-acetyl trypsin and incubated at 37 °C for 48 h. Cells were fixed in 5% formaldehyde
for 1 h at room temperature. Plaques were visualized by crystal violet staining. Adeno-associated virus was prepared according to McClure _et al._ 57 .

Reagents. 2% w/v methylcellulose (25 centipoise, Sigma Aldrich, Dorset, UK) was prepared as described
in Griffiths _et al._ 27 . Gold particles were prepared by citrate reduction according to Frens 44 and stored at 4 °C. To
make the calibration solution, freshly made colloid was mixed 1:1 with 2% methylcellulose. To remove citrate
and sodium ions, the mix was centrifuged for 2 hours at 8000 × g at 4 °C and sedimented particles resuspended
in fresh 1% methylcellulose. The calibration solution was stored at months at 4 °C. Nanodiscs were prepared
as described 35, and liposomes according to Marius _et al._ 41 . Standard plastic support films were made using 1%

pioloform (Agar Scientific, Stansted, Essex, UK) and copper hexagonal 100-mesh or square 200 mesh grids
applied to the film. Nanorods (10 nm thick) were obtained from Sigma Aldrich (Dorset, UK).

Stain mixes and application to EM grids. Basic nanoparticle staining mixture (without incorporating
gold calibration or pre-dialysis) contained 1.5 μ l of 1% methylcellulose mixed with 7.5 μ l of nanoparticle solution
and 1 μ l of 0.3% uranyl acetate (values are given in μ l to illustrate minimal volumes but larger volumes were also
used in proportion; final uranyl acetate and methylcellulose concentrations: 0.03% and 0.15% respectively). When
calibration was required, 1.5 volumes of gold particle-calibration solution (containing 1% methylcellulose) were
added to 1.5 volumes of nanoparticle solution, 1 volume of 0.3% uranyl acetate and 6 volumes of deionized water.
When dialysis was required, equal volumes of calibration solution and nanoparticle solution were mixed and
capillary dialysis carried out as detailed below. 0.5 μ l of staining-mix were loaded onto pioloform-coated EM grids
using a standard 2 μ l micropipette under binocular control. The grids were then air-dried and examined in a JEOL
1200 EX transmission electron microscope operated at 80 kV and imaging carried out using a GATAN Orius 200
digital camera (GATAN, Abingdon, Oxon, UK).

Capillary tube dialysis. Renal hemodialysis cassettes contain numerous parallel capillary straws that were
best extracted by pulling groups from the bundle, using forceps. Immediately after drawing them out from the
cassette, each straw was cut with a clean scalpel at each end. Then each end was passed through the narrower
aperture of a truncated micropipette tip (as shown in Fig. 3) and the mix aspirated by capillary action. To seal
the tube, another pipette tip was then inserted into the wider aperture of the first tip, thereby trapping the capillary. The whole assembly was subsequently immersed in dialyzing solution. After dialysis the tube was laid onto
parafilm and curved forceps drawn along the tube to expel the dialysate. The drop was retrieved and stored in an
Eppendorf centrifuge tube at 4 °C.

Counting and Sizing procedures. These procedures were carried out directly on a digital camera display
(see Figs 4 and 5). Counting was done by manual translocation of the specimen in a single direction and particles selected using unbiased counting rules applied to the edges of a counting band. The edges of the band were
defined by lines traced out by two arrows positioned inside opposite sides of the image frame, thereby creating
a peripheral guard area. This allowed morphological characterization/size estimation even of the largest nanoparticles. During scanning one edge of one of the spots defined an acceptance line and the edge of the other
spot defined a forbidden line. Any particle that fell completely within the lines, or was crossed by the acceptance
line, was sampled/accepted for counting. Any particle that intersected the forbidden line was not sampled or
counted 42,43 .

To evaluate the influence of methylcellulose on the binding of nanoparticles to the support film, liposomes
were mixed 1:1 with 2% methylcellulose or deionized water and the mix allowed to adsorb onto an EM grid
for 2 minutes. After three washes on droplets of deionized water (2 minutes per step), followed by contrasting
with 3% (w/v) uranyl acetate/2% methylcellulose (ratio of 1:9); grids were air-dried in a wire loop (according
to Griffiths _et al._ 27 ). The number of liposomes per unit area was assessed in micrographs taken SUR at 2,000× .
Counting of liposomes was carried out in rectangular quadrats measuring 2.1 μ m × 2.1 μ m to which unbiased
counting rules were applied 43 and counts related to the area.

For estimating particle caliper diameter, a systematic series of dots with regular spacing was placed orthogonal to the travel direction of the microscope stage. The spacing was set to about half the size of the smallest
nanoparticle and the extent of the array was greater than the width of the counting band so that any peripheral
particles sampled by the counting band were sized. Consistently, one extreme edge of each dot was assigned for
counting intersections and the number of edges that crossed each particle during translocation of the specimen
was counted. An unbiased estimate of the particle caliper distance is ∑ I × d where I is the number of intersections
and d the real spacing between the dots.

To compare a random point process to the distribution of calibration nanoparticles, 20 micrographs, positioned systematically with a random start (SUR), were taken for each nanoparticle (magnification 3000 x for
liposomes and 4000 x for influenza A virus). The images were opened in Photoshop CS6 and gold particle counts
made using the unbiased counting rule described above (quadrat areas 9.49 μ m 2 for liposomes and 6.15 μ m 2 for
influenza A virus). The gold calibration particles were categorized as “associated” or “non-associated” with nanoparticles (particles less than one particle width distant from the nanoparticle were classified as associated). For
the distribution of random points the same micrographs were overlaid with a randomly placed square systematic
grid lattice (spacing 2.05 μ m for liposomes and 1.55 μ m influenza A virus) resulting in 1 to 4 random points per
placement. The random points were categorized as for the calibration particles. The process of random grid placement was repeated until the number of random points matched gold particles for the corresponding micrograph.

To assess the relationship between estimated nanoparticle number and dilution, a dilution series of liposomes
was prepared in deionized water using 10 μ l of neat liposome solution for every dilution (1/50, 1/100, 1/200 and
1/1000). For each sample four staining mixes were prepared and 0.5 μ l per mix loaded onto individual EM grids.
The grids were then quantified and the liposome number for each sample estimated (see above) and plotted in
Microsoft Excel.

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Acknowledgements
We thank Dana Hassan and Benjamin Griffin for testing the method as untrained participants. We are grateful
to Luis Cruz-Orive for assistance with calculations of profile width after projection from a tilted oblate spheroid.
This work was supported by the University of St Andrews and Nanomorphomics group funds. The work forms
part of an International Patent Application No. PCT/GB2015/052482. This manuscript is dedicated to the
memory of Kiyoteru Tokuyasu.

Author Contributions
J.M.L. conceived, designed and coordinated/supervised the experiments, wrote the manuscript and performed
calibration and nanorod experiments. C.H. performed experiments, optimized methods, prepared manuscript
figures and contributed to writing the methods. J.A. performed initial pilot studies. C.P. prepared all nanodisc
samples and MscL containing liposomes. S.F. performed nanoparticle quantification experiments and edited
the manuscript and figures. P.M. prepared liposomes and J.T. prepared adeno-associated virus. J.H.N. and D.J.
analyzed data and contributed to writing the paper. L.S. prepared the influenza A virus and performed the plaque
forming unit-assays.

Additional Information
Supplementary information [accompanies this paper at http://www.nature.com/srep](http://www.nature.com/srep)

Competing financial interests: The authors declare no competing financial interests.

How to cite this article : Hacker, C. _et al._ Nanoparticle suspensions enclosed in methylcellulose: a new approach
for quantifying nanoparticles in transmission electron microscopy. _Sci. Rep._ 6, 25275; doi: 10.1038/srep25275
(2016).

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