Insight into cell-entry mechanisms of CPPs by electron microscopy.

Abstract

Despite the quickly widening application of cell-penetrating peptides (CPP) for the cellular delivery of various macromolecules, the cell entry mechanisms of these peptides have remained elusive so far. The basic features of the translocation of CPPs into cells have been mapped by fluorescence microscopy and activity-based assays revealing that endocytotic mechanisms are mainly responsible for the uptake at physiological temperature. However, the high concentration of CPP or the lowering of the incubation temperature below 10°C (re)activates a nonvesicular cell entry mode. The fluorescence microscopy can hardly provide detailed information about the interaction of CPP molecules with the extracellular structures, the induced changes in the morphology of the plasma membrane, etc. Therefore, application of electron microscopy could help to shed light on the nature of nonvesicular uptake mechanism. Transmission electron microscopy (TEM) has been a valuable tool for the morphological characterization of biological material at high resolution. It can provide useful information at the ultrastructural level about the interaction and arrangement of CPPs on the cell surface, the entrapment in cellular organelles and the translocation to the cytoplasm. In this chapter, we present a method for the tagging of CPPs covalently with a 1.4 nm gold cluster and provide a flat-embedding protocol for the mapping of Nanogold™-labeled CPPs in cultured cells by TEM. This method enables to retain the cell monolayers in their in situ orientation. The Nanogold™ tag is putatively not interfering with the uptake of CPPs and enables the production of specimens with excellent morphology and good contrast.

Full text

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Chapter 13
Insight into Cell-Entry Mechanisms of CPPs
by Electron Microscopy
Kärt Padari, Annely Lorents, Eija Jokitalo, and Margus Pooga
Abstract
Despite the quickly widening application of cell-penetrating peptides (CPP) for the cellular delivery of
various macromolecules, the cell entry mechanisms of these peptides have remained elusive so far. The
basic features of the translocation of CPPs into cells have been mapped by fluorescence microscopy and
activity-based assays revealing that endocytotic mechanisms are mainly responsible for the uptake at
physiological temperature. However, the high concentration of CPP or the lowering of the incubation
temperature below 10°C (re)activates a nonvesicular cell entry mode. The fluorescence microscopy can
hardly provide detailed information about the interaction of CPP molecules with the extracellular struc-
tures, the induced changes in the morphology of the plasma membrane, etc. Therefore, application of
electron microscopy could help to shed light on the nature of nonvesicular uptake mechanism.
Transmission electron microscopy (TEM) has been a valuable tool for the morphological characteriza-
tion of biological material at high resolution. It can provide useful information at the ultrastructural level
about the interaction and arrangement of CPPs on the cell surface, the entrapment in cellular organelles
and the translocation to the cytoplasm. In this chapter, we present a method for the tagging of CPPs
covalently with a 1.4 nm gold cluster and provide a flat-embedding protocol for the mapping of
Nanogold™-labeled CPPs in cultured cells by TEM. This method enables to retain the cell monolayers
in their in situ orientation. The Nanogold™ tag is putatively not interfering with the uptake of CPPs and
enables the production of specimens with excellent morphology and good contrast.
Key words: Cell-penetrating peptide, Transmission electron microscopy, Nanogold-labeled CPP
CPPs have been used as efficient carriers in drug and gene delivery
enabling specific targeting and resulting in high biological
response (1–3). However, the current knowledge about how
CPPs reach their target compartments inside cells is still far from
complete. To better understand the uptake mechanism and distri-
bution of CPPs and their cargo molecules, it is essential to apply
1. Introduction
Ülo Langel (ed.), Cell-Penetrating Peptides: Methods and Protocols, Methods in Molecular Biology, vol. 683,
DOI 10.1007/978-1-60761-919-2_13, © Springer Science+Business Media, LLC 2011

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182Padari et al.
different complementary methods in parallel, which would allow
adequate interpretation of experimental data (4). Most of the
studies examining the uptake and cellular localization of CPPs
have used fluorescence microscopy. Even though fluorescence
microscopy is an excellent tool for real-time studies in living cells
and can be successfully used for quantitative analysis, it has its
limitations. First, the fluorescence signal might be quenched in
the cellular environment, especially in the vesicles with low pH or
when associated with polyanions, as well as upon interaction with
cell surface proteoglycans (5), complicating the interpretation of
fluorescence microscopy results. Second, due to the limited reso-
lution of light microscope, it cannot provide detailed information
about the interaction of CPPs with the plasma membrane and
their exact localization in relation to intracellular compartments.
However, detailed information can be acquired by transmission
electron microscopy (TEM) that allows visualization of subcellu-
lar compartments at the ultrastructural level to study the associa-
tion with the membranes, the uptake, and the intracellular
trafficking of CPPs with very high precision. Still, relatively few
studies have applied electron microscopy for assessing the uptake
mechanisms of CPPs. The main drawback is the need to fix the
specimen that can lead to artifactual redistribution of some peptides
into, but also inside of cells (6). Still, not all CPPs redistribute in
cells upon fixation (7) and the localization of CPP–cargo conjugates
is influenced even less by treatment with fixatives (8), making
TEM studies in this field feasible and justified.
TEM has mostly been used in studies focused on the CPP-
mediated delivery of proteins (9–11) or gold nanoparticles alone
(12–14) rather than for the characterization of internalization
mechanisms and intracellular fate of CPPs themselves. For exam-
ple, the translocation of 16 nm gold nanoparticles modified with
Tat-peptide and Penetratin into cell interior was characterized
recently and the particles were found in endosomes along with a
dispersed signal in the cytosol (12). However, considering that the
size of the cargo molecule might determine the uptake mechanism
of CPPs (15, 16), a small nanogold tag (1.4 nm) rather than col-
loidal gold particles (14–16 nm) have to be harnessed in the stud-
ies of CPP mechanisms. In addition, the Nanogold™ (NG) cluster
is coupled to the CPP molecule by a covalent bond, resulting in
homogenous well-defined compound on the contrary to colloidal
gold, in which neither the composition of label nor the number of
peptide molecules per particle can be exactly defined.
A recent study revealed that the novel CPPs derived from
perforin and granzyme do not associate with the cell surface ran-
domly but assemble in spherical structures. The clusters of
NG-labeled CPP interfered with the regular packing of the lipid
bilayer as the plasma membrane became less distinct in TEM.
Although the novel CPPs are taken up by cells mostly by endocytotic

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Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
mechanisms, the peptide clusters are not dissociated in the hostile
milieu of the endosomes during the first hours (17, 18).
In this chapter, we will discuss the method of TEM for map-
ping the interaction of NG-labeled CPPs with cells and the fol-
lowing uptake. We present the protocols for the labeling of CPPs
with preactivated nanogold tag and embedding the cultured cells
in a resin by so-called flat-embedding technique, which retains
the orientation and morphology of cells.
In this chapter, we describe protocols for cultivating HeLa cells
for TEM. HeLa cells are derived from the human cervical carci-
noma and have been used in a significant number of other CPP
studies. Using the same cell line enables one to complement and
compare experimental data with other research groups in the
field. However, any other cell line or primary cells can also be
used in the studies. Still, for the flat-embedding protocol pro-
vided here, only adherent cells can be used.
1. CPP with a thiol group (see Note 1).
2. Monomaleimido nanogold (Nanoprobes Inc., Yaphank, NY)
or monomaleimido undecagold.
3. Oxygen-free MilliQ water. Remove air dissolved in MilliQ
water by vacuum followed by bubbling through argon for at
least 15 min.
4. 50% Methanol (³99.9%) in oxygen-free water.
5. Oxygen-free MilliQ water with 0.1% trifluoroacetic acid
(TFA ³ 99.9%) (see Note 2).
6. Acetonitrile with 0.1% TFA.
1. Eppendorf tubes filled with argon.
2. Spectrophotometer (e.g., Nanodrop 1000, Thermo Fischer
Scientific Inc.).
3. Thermostat mixer (e.g., Thermomixer Comfort, Eppendorf
AB, Germany).
4. Rotational vacuum concentrator (e.g., RVC 2-25, Christ
GmbH, Germany, or Savant Speed-Vac SC110, Ramsey,
MN).
5. Chromatography system equipped with columns for peptide
purification and gel filtration by reversed phase chromatog-
raphy (see Note 3).
2. Materials
2.1. Cell Culture
2.2. Labeling
of Peptide
2.2.1. Reagents
and Solutions
2.2.2. Materials
and Equipment

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184Padari et al.
1. Human cervical carcinoma cell line HeLa cultured in Iscove’s
Modified Dulbecco’s Medium (IMDM) supplemented with
10% fetal bovine serum (FBS), 100 IU/mL penicillin, and
100 mg/mL streptomycin.
2. IMDM supplemented with 100 IU/ml penicillin, 100 mg/ml
streptomycin, and 10% FBS for the incubation of cells with
CPP–nanogold (CPP–NG) conjugates.
3. Sodium cacodylate buffer: prepare 0.4 M stock solution by
dissolving 21.4 g of cacodylic acid sodium salt trihydrate
[Na(CH3)2AsO2⋅3H2O] in 250 ml MilliQ water (see Note 4).
Adjust pH to 7.4 by adding 0.2 M HCl (about 8 ml) to 50 ml
of stock solution and add MilliQ water to the volume of
200 ml to make a 0.1 M working solution. Alternatively, to
make 0.2 M working solution (for preparing osmium tetrox-
ide solution, see step 9 below) dilute to final volume of
100 ml.
4. Fixative: 2.5% glutaraldehyde in 0.1 M cacodylate buffer, pH
7.4. Always use freshly prepared fixative. Store 25% glutaral-
dehyde stock [Electron Microscopy Sciences (EMS), Hatfield,
PA] in aliquots at −20°C and do not use the reagent that has
been thawed and frozen again.
5. Silver enhancement reagent: HQ Silver Kit (Nanoprobes Inc.,
Yaphank, NY). The components of HQ Silver kit can be ali-
quoted in amber Eppendorf tubes and stored at −20°C. The
components of silver enhancement kit are light sensitive,
therefore it is necessary to work in dark room under red safe
light conditions while aliquoting or staining specimens.
6. Reagents for gold toning:
(a) 2% sodium acetate [CH3COONa⋅3H2O] in MilliQ water
(use freshly made solution).
(b) 0.05% gold chloride [HAuCl4⋅H2O] in MilliQ water (can
be stored at 4°C for several months).
(c) 0.3% sodium thiosulfate pentahydrate [Na2S2O3⋅5H2O]
in MilliQ water (use freshly prepared solution).
7. Reduced, buffered osmium tetroxide solution for the staining
and postfixation of the specimens (see Note 5). Mix equal
volumes of 2% OsO4 in MilliQ water and 0.2 M cacodylate
buffer to yield a 1% OsO4 in 0.1 M cacodylate buffer. Add
15 mg potassium ferrocyanide (K4[Fe(CN)6]) per 1 ml of 1%
OsO4 solution (see Note 6).
8. 70% Ethanol in MilliQ water.
9. 96% Ethanol in MilliQ water.
10. Ethanol (³99.5%).
11. Acetone (³99.5%).
2.3. Preparation
of a Specimen for TEM

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Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
12. Embedding resin: TAAB Premix Embedding Kit (medium,
TAAB Laboratories Equipment Ltd, UK) (see Note 7).
13. 2% Uranyl acetate (UA) in 50% ethanol (see Note 8).
14. Lead citrate stain: Add 20 mg of lead citrate [Pb3(C6H5O7)2⋅
3H2O] to 10 ml of CO2-free MilliQ water (boil water for
10 min to make it CO2 free) (see Note 9). Add 0.1 ml of
10 N NaOH (see Note 10), seal the tube air-tightly and shake
vigorously until all the lead citrate is dissolved. Filter the solu-
tion through a 0.2 mm Millipore filter before use.
15. Liquid nitrogen for removing coverslips from polymerized
resin blocks of specimen.
16. Single-use plastic Pasteur pipettes (transfer pipettes) for
changing solutions.
17. Parafilm.
18. Embedding capsules and capsule holder: We use BEEM®
embedding capsules (size 3) and BEEM® capsule holder from
EMS. However, any other suitable capsules and holders can
be used.
19. Aluminum planchettes or dishes.
20. 24-Well culture plates.
21. Round glass coverslips (Ø 12 mm, no. 1).
22. Cell culture dishes (35 × 10 mm).
1. Dissolve the lyophilized peptide in oxygen-free MilliQ water
to yield a 0.5 mM peptide solution. Calculate the volume of
water for dissolving the peptide batch based on its weight and
molecular mass (see Note 11).
2. Check the concentration of peptide by measuring the optical
density of the prepared solution using a suitable dilution, at
280 nm if the peptide contains tryptophan or tyrosine (see
Note 12) or at 210 nm. If the weight-based concentration
differs from absorbance-based concentration by more than
10%, use the latter.
3. Dissolve monomaleimido nanogold in 50% methanol at
30 mM concentration (see Note 13).
4. Couple the label to peptide by adding CPP solution (2.5-fold
molar excess) in small aliquots to nanogold solution upon
stirring (see Note 14). Incubate the mixture for 60–90 min at
30°C in dark under mild stirring.
3. Methods
3.1. Labeling
of Peptides with
Nanogold

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186Padari et al.
5. Remove methanol and concentrate the solution of conjugate
by rotational vacuum concentrator at 30°C to reach ~100 mM
concentration (see Note 15).
6. Measure the absorbance of the resulting solution at 280 nm
using the suitable dilution and calculate the concentration of
conjugate considering that 1 A280/cm corresponds to ~2.1 mM
nanogold (see Note 16). Aliquot the conjugate and store in
freezer. Store the working aliquot in fridge and use it within
1 month (some conjugates can be stored in fridge less than 1
week). Avoid the repeated freeze–thaw cycles.
7. Purify the conjugate by reversed phase chromatography or
gel filtration if necessary (see Note 17).
8. Concentrate the fractions of chromatographic purification by
rotational vacuum concentrator (see step 5), pool if necessary
and measure the concentration of the conjugate (see Note 15).
1. Clean glass coverslips to remove any dust particles by rinsing
multiple (3 to 4) times with 70% ethanol in a 100 ml glass
flask. Dry and sterilize coverslips one by one in the flame of
gas burner using tweezers.
2. Place the sterilized coverslips to the bottom of cell culture
dish (35 × 10 mm). We recommend having at least three cov-
erslips in one culture dish to get three parallels of each set of
experiment for different time points of silver-enhancement
procedure (see Subheading 3.3 below). You can also use up
to four coverslips (i.e., four different time points of silver
enhancement) per one set of experiment.
3. Seed HeLa cells onto coverslips in culture dish and grow for
2 days to reach 80–100% of confluence (see Note 18).
4. Remove the culture medium and incubate the cells with
CPP–NG conjugates in IMDM (1 ml solution per culture
dish with three coverslips) at desired concentration and tem-
perature for required time depending on your experiment
(for example 1 mM of CPP–NG at 37°C for 1 h).
5. Take cells out from the incubator just before the fixation and
wash twice with prewarmed IMDM.
6. Remove the medium and immediately apply the fixative.
Make sure that the cells do not dry at any stage during the
procedures. Fix the cells with 2.5% glutaraldehyde in cacody-
late buffer for 30–60 min at room temperature (RT).
7. Wash with cacodylate buffer for three times for 10 min (here
you can interrupt the procedure and leave samples in buffer
for overnight at 4°C). Continue with silver enhancement (see
Subheading 3.3).
3.2. Treatment of Cells
with CPP–NG
Conjugates and
Fixation

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Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
For most applications, the detection of nanogold particles in an
electron microscope specimen requires the enlargement of particles
in order to visualize these with the magnification range routinely
used for imaging cellular organelles. Gold particles act as catalysts
in the presence of silver ions and a reducing agent and reduce silver
ions to metallic silver. The silver is deposited onto the gold enlarg-
ing thereby the 1.4 nm gold particles to about 10 nm diameter or
more depending on the enhancement time and temperature. Silver
enhancement reaction is time dependent: nanogold particles
enlarge particularly rapidly within the first minutes, later the growth
rate declines since the surface area of particles increases. After the
suggested time of the enhancement, silver may precipitate sponta-
neously by self-nucleation yielding a background signal. Therefore,
it is very important to choose the optimal time for enhancement in
order to magnify the nanogold particle without increasing the
background staining. Additionally, the deposition of silver onto the
gold surface is quicker at higher temperatures. Therefore, we usu-
ally prepare two to three specimens in parallel using different dura-
tion of silver enhancement. It is particularly important for the
protocol provided here because the enhancement is carried out
before embedding and if the enhancement proves to be too long
(resulting in too large particles) it cannot be reversed.
The following steps of the silver enhancement and gold ton-
ing procedures should be carried out in a dark room under red
light at 20–22°C.
1. Wash the cells three times with MilliQ water for 3 min.
2. Prepare a 24-well plate for stopping enhancement reaction
(see step 5) by filling wells with MilliQ water.
3. Mix equal amounts (for example 200 ml) of the initiator,
moderator, and activator just before use (see instructions of
HQ SILVER Enhancement Kit).
4. Place three drops (about 50 ml) of enhancing solution on para-
film and place coverslips upside down onto the drops to enlarge
the particles for 1, 3, and 5 min. If you have four parallels,
incubate coverslips for 30 s, 1, 2, and 4 min. Be as precise as
possible with incubation times (use a timer) (see Note 19).
5. Stop the reaction by transferring coverslips (cells upside) to
MilliQ water in wells of culture plate.
6. Wash twice with MilliQ water for 5 min.
7. Stabilize the silver-deposited particles by gold toning (see
Note 20):
(a) Wash three times with 2% sodium acetate for 5 min at RT.
(b) Treat with 0.05% gold chloride for 10 min on ice.
(c) Wash twice with 0.3% sodium thiosulfate (freshly made)
for 10 min on ice.
3.3. Silver
Enhancement

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188Padari et al.
8. Wash three times with MilliQ water for 3 min.
9. Continue with osmication (see Subheading 3.4).
1. Osmicate with 1% OsO4 for 1 h at RT. Avoid direct daylight
by covering the culture plate with aluminum foil during
osmication.
2. Wash three times with 0.1 M cacodylate buffer for 5 min.
3. Dehydrate once with 70%, once with 96% and twice with
absolute ethanol for 1 min each step.
4. Remove the caps from the embedding capsules, place the
capsules in holder and fill with embedding resin to maximum.
Keep in mind to take resin (if stored in syringes at −20°C) out
from freezer before the embedding procedure and let them
warm up to RT before use.
5. Dip a coverslip into acetone for couple of seconds and place
it onto an aluminum planchette (cells upside).
6. Drop immediately some embedding resin to the cells. Do not
allow the complete evaporation of acetone.
7. Place the capsules filled with embedding resin upside down
on the top of cells.
8. Keep specimens at RT for 2 h to allow the resin to infiltrate
into cells.
9. Transfer the cells to 60°C oven and polymerize overnight (for
at least 14 h).
10. After the polymerization, remove the coverslips by dropping
samples directly from the oven to liquid nitrogen and crack
the coverslip from the block. Make sure that no pieces of glass
remain on the resin block; otherwise these could damage the
cutting edge of your diamond knife during sectioning (see
Note 21).
11. Cut cells embedded in resin into ultrathin sections (30–
50 nm, i.e., silver gray sections) and collect on the copper
grids (EMS).
12. Stain the sections with 2% UA on parafilm for 1 min.
13. Wash twice in 50% ethanol and let the sections dry in air for
15–20 min.
14. Wash the sections with 0.01 N NaOH for 1 min.
15. Stain the sections with solution of lead citrate for 1–3 min
depending on the required contrast.
16. Wash with 0.01 N NaOH and rinse thoroughly (at least three
times) by transferring grids from one drop of MilliQ water to
another.
17. Let the grids dry and examine in transmission electron micro-
scope operated at 80 kV.
3.4. Embedding of
Cells (Flat-Embedding)

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Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
1. Peptides could also be tagged with nanogold on the amino
group of lysine or N terminus, but such modification decreases
the net positive charge and usually reduces the cellular uptake of
CPPs. Therefore the cysteine residue is introduced in CPPs.
2. MilliQ water and acetonitrile with TFA are necessary only if
the CPP–nanogold conjugate is purified by reversed phase or
gel-filtration (size exclusion) chromatography.
3. In this work, an FPLC system (GE Healthcare/Pharmacia)
equipped with absorbance detectors at 280 and 210 nm, an
automated fraction collector and the columns Superdex-
Peptide HR10/30 (for gel filtration chromatography) and
Pro RPC HR5/2 (for reversed phase chromatography)
were used.
4. Sodium cacodylate contains arsenic, which is a health hazard
if inhaled or absorbed through the skin. Use gloves and fume
hood for weighing the reagent and preparing the buffer solu-
tion. Do not let the reagent come in contact with acids in
order to avoid the production of arsenic gas (19).
5. Osmium tetroxide must be handled with the utmost care
because of its high toxicity. Exposure to OsO4 vapor can cause
severe eye, skin, and respiratory problems. Prepare the
osmium tetroxide solutions always under the vented hood
and handle bottles with disposable gloves. To make 2% aque-
ous solution of OsO4, we use the OsO4 crystals in the glass
ampoule (available from EMS). We recommend immersing
the ampoule into liquid nitrogen prior the opening in order
to crystallize vaporized osmium tetroxide. OsO4 is also sup-
plied as an aqueous solution in glass ampoules. Store the
solution of 2% OsO4 at 4°C in a clean brown glass bottle to
avoid the contamination by organic matter and exposure to
light. Since the vapors of osmium tetroxide can readily leak
out of many containers, use double glass bottles sealed with
parafilm for storing OsO4. All used OsO4 solutions should be
collected into a glass bottle containing vegetable oil (corn oil
is preferred because of its high percentage of unsaturated
bonds) and stored in the fume hood. For full neutralization,
two volumes of unsaturated oil per one volume of 2% osmium
tetroxide solution are needed. The neutralized osmium solu-
tion is then disposed in accordance to each country’s regula-
tions. Contact your environmental health and safety office to
obtain local regulation.
6. Osmium tetroxide in combination with ferrocyanide is used
for enhancing the contrast of many cellular components,
including membranes and glycogen.
4. Notes

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190Padari et al.
7. Components of the embedding kit can be stored at 4°C for
12 months. The mixed and ready-to-use embedding resin can
be stored, e.g., in plastic syringes without needle (5 or 2.5 ml)
at −20°C for later use for 3 months. Caps for syringes are
available from EMS. Most embedding resins are carcinogenic!
Cover working area with paper before mixing the compo-
nents and always use disposable gloves during embedding
procedures. Seal used tubes, pipettes, dishes, etc. in a plastic
bag and polymerize all resin waste before disposal at 60°C for
overnight. Never pour any resin containing solutions down
the drain, the resin will polymerize and could clog the tubes.
8. Centrifuge UA solution before use. Care should be taken in
the handling and disposal of uranium-containing solutions
because of their radioactivity and chemical toxicity. Danger of
cumulative effects!
9. Use only carbonate-free MilliQ water and freshly made NaOH
to prevent the formation of lead carbonate precipitate. To avoid
contamination of the staining solution with carbonate ions,
keep the tube tightly sealed from atmospheric carbon dioxide.
Use the staining solution within the day of preparation.
10. Prepare 10 N sodium hydroxide by dissolving 4 g of NaOH
in 10 ml of CO2-free distilled water. Be aware that dissolution
of sodium hydroxide is highly exothermic.
11. The peptides with a thiol group oxidize very easily to form
dimers, therefore, for the labeling procedure do not use the
stock solution of peptide, which has been stored for a long
time, or is prepared in water that contains oxygen. CPPs can
be dissolved at any desired concentration. However, at very
high concentration more hydrophobic peptides tend to pre-
cipitate upon labeling and low peptide concentrations might
result in low labeling yield.
12. Calculate the molar extinction coefficient of your peptide as
follows: molar extinction coefficient = (number of tryptophan
residues × 5,500) + (number of tyrosine residues × 1,490)
(http:/ /www.encorbio.com/protocols/Prot-MW-Abs.htm).
13. Reconstitution of 30 nmol batch of monomaleimido nano-
gold with 1 ml deionized water will yield a 20 mM sodium
phosphate buffer at pH 6.5 with 150 mM NaCl, i.e., suitable
conditions for selective labeling of thiol groups of peptide.
Methanol (or acetonitrile) facilitate the dissolving of the label
and reduce precipitation of hydrophobic CPPs during label-
ing. However, monomaleimido nanogold can be reconsti-
tuted in a smaller or a larger volume of solvent if necessary.
14. Perform the coupling reaction in a tube filled with argon to
maximize the yield of coupling by excluding the oxidation of
thiols. Use a two- to fourfold molar excess of CPP over the

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Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
monomaleimide nanogold to assure that most of the label
reacts with the peptide. Addition of the CPP solution in water
to nanogold solution in 50% methanol decreases the possibil-
ity of the peptide/conjugate precipitation during labeling.
15. Check the volume of the conjugate solution during the con-
centration step regularly (every 30–60 min, depending on the
initial volume). Some amphipathic/hydrophobic leucine-rich
CPPs may precipitate during methanol removal and concen-
tration. This could be decreased or even avoided by adding
some concentrated stock solution of leucine to the reaction
mixture before concentration step (final concentration
2–5 mg/ml depending on CPP).
16. Tryptophan contributes to the absorption of a conjugate at
280 nm by 1% and may therefore not be taken into account.
Typically >90% of nanogold is coupled to the peptide under
the used reaction conditions, and the concentration of the
conjugate can be calculated based on the total absorption of
the solution at 280 nm.
17. The purification of the conjugate to homogeneity is often not
necessary and the mixture with unlabeled peptide (the con-
centrated reaction mixture) can be used in the majority of
cellular localization studies. The inactive (hydrolyzed)
monomaleimide nanogold (in analogy with cationized or
neutral NG) neither strongly binds to the extracellular matrix
nor is taken up by cells via endocytosis. The unlabeled CPP
remains invisible in TEM and does not interfere with the
analysis of the subcellular localization of CPP–NG conju-
gates. Most importantly, due to their highly cationic nature,
CPPs adhere strongly to various surfaces, which lead to high
losses of peptides and their conjugates with NG in purifica-
tions by chromatographic methods. Run the reversed phase
chromatography in C4 to C8 column of minimal necessary
volume. We used Pro RPC HR5/2 (Pharmacia, Sweden) and
run a steep gradient from 10% acetonitrile in water (both with
0.1%, v/v TFA) to 100% acetonitrile. The CPP–NG conju-
gate elutes in about 10% higher acetonitrile concentrations
than the peptide. Different CPPs elute from the column in
different conditions, therefore, optimize the shape of the gra-
dient with a minimal amount of the conjugate before the
preparative purification. Gel-filtration chromatography is sug-
gested for the purification of nanogold-labeled proteins and
peptides by Nanoprobes Inc. However, the gel filtration
chromatography does not separate well the hydrolyzed NG
label from the conjugate with CPP due to the small difference
in size. Moreover, the conjugates of CPP with NG could be
recovered in very low yield (for Superdex peptide HR 10/30
<50%). Addition of a volatile solvent (e.g., acetonitrile up to

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192Padari et al.
40%) reduces the losses in purification to some extent, but
the recovery yield still remains poor.
18. For electron microscopy, the cells have to be grown on glass
coverslips for at least 2 days to guarantee stable adhesion and
good morphology of cells in specimens.
19. We have used HQ SILVER™ Enhancement kit (Nanoprobes)
in our studies, which enables controlled silver enhancement
and yields homogenous particles. However, it should be kept
in mind that the adjacent particles might fuse and form irreg-
ular shapes upon growth. Therefore very long incubations
that result in large particles should be avoided.
20. Gold toning is the posttreatment of silver enhanced gold par-
ticles with gold chloride. This treatment deposits a thin layer
of gold onto the surface of the particles and stabilizes the
silver deposition, making the resulting particles of Au–Ag–Au
more resistant to the following treatments with osmium tet-
roxide and UA solutions (20).
21. Allow blocks to settle at RT for couple of hours before trim-
ming a small pyramid suitable for successful ultrathin section-
ing. Small block face facilitates the parallel alignment of block
face and knife edge that is important as the specimen thick-
ness is only one cell layer thick thus allowing no trimming
from the block face.
Acknowledgments
We thank the people of Electron Microscopy Unit at the University
of Helsinki for introducing the silver enhancement method and
improving the flat-embedding technique; and M. Kure for excel-
lent technical assistance in electron microscopy. The work was
supported by grants from Estonian Science Foundation (ESF
7058) and Estonian Ministry of Education and Research
(0182691s05).
References
1. El-Andaloussi, S., Johansson, H., Lundberg,
P., and Langel, Ü. (2006) Induction of splice
correction by cell-penetrating peptide nucleic
acids. J Gene Med 8, 1262–73.
2. Wu, B., Moulton, H. M., Iversen, P. L.,
Jiang, J., Li, J., Spurney, C. F., Sali, A.,
Guerron, A. D., Nagaraju, K., Doran, T., Lu,
P., Xiao, X., and Lu, Q. L. (2008) Effective
rescue of dystrophin improves cardiac function
in dystrophin-deficient mice by a modified
morpholino oligomer. Proc Natl Acad Sci
USA 105, 14814–9.
3. Mäe, M., El-Andaloussi, S., Lundin, P.,
Oskolkov, N., Johansson, H. J., Guterstam,
P., and Langel, Ü. (2009) A stearylated CPP
for delivery of splice correcting oligonucle-
otides using a non-covalent co-incubation
strategy. J Control Release 134, 221–7.

Page 13

193
Insight into Cell-Entry Mechanisms of CPPs by Electron Microscopy
4. Holm, T., Johansson, H., Lundberg, P.,
Pooga, M., Lindgren, M., and Langel, Ü.
(2006) Studying the uptake of cell- penetrating
peptides. Nat Protoc 1, 1001-5.
5. Ziegler, A., and Seelig, J. (2007) High affinity
of the cell-penetrating peptide HIV-1 Tat-
PTD for DNA. Biochemistry 46, 8138–45.
6. Lundberg, M., and Johansson, M. (2001) Is
VP22 nuclear homing an artifact? Nat
Biotechnol 19, 713–4.
7. Mano, M., Teodosio, C., Paiva, A., Simoes,
S., and Pedroso de Lima, M. C. (2005) On
the mechanisms of the internalization of
S4(13)-PV cell-penetrating peptide. Biochem
J 390, 603–12.
8. Säälik, P., Elmquist, A., Hansen, M., Padari,
K., Saar, K., Viht, K., Langel, Ü., and Pooga,
M. (2004) Protein cargo delivery properties
of cell-penetrating peptides. A comparative
study. Bioconjug Chem 15, 1246–53.
9. Padari, K., Säälik, P., Hansen, M., Koppel, K.,
Raid, R., Langel, Ü., and Pooga, M. (2005)
Cell transduction pathways of transportans.
Bioconjug Chem 16, 1399–410.
10. Magzoub, M., Sandgren, S., Lundberg, P.,
Oglecka, K., Lilja, J., Wittrup, A., Goran
Eriksson, L. E., Langel, Ü., Belting, M., and
Gräslund, A. (2006) N-terminal peptides
from unprocessed prion proteins enter cells by
macropinocytosis.
Commun 348, 379–85.
11. Säälik, P., Padari, K., Niinep, A., Lorents, A.,
Hansen, M., Jokitalo, E., Langel, Ü., and
Pooga, M. (2009) Protein delivery with trans-
portans is mediated by caveolae rather than
flotillin-dependent pathways. Bioconjug Chem
20, 877–87.
12. Nativo, P., Prior, I. A., and Brust, M. (2008)
Uptake and intracellular fate of surface-modi-
fied gold nanoparticles. ACS Nano 2,
1639–44.
Biochem Biophys Res
13. Pujals, S., Bastus, N. G., Pereiro, E.,
Lopez-Iglesias, C., Puntes, V. F., Kogan, M.
J., and Giralt, E. (2009) Shuttling gold nano-
particles into tumoral cells with an amphip-
athic proline-rich peptide. Chembiochem 10,
1025–31.
14. Mandal, D., Maran, A., Yaszemski, M. J.,
Bolander, M. E., and Sarkar, G. (2009) Cellular
uptake of gold nanoparticles directly cross-
linked with carrier peptides by osteosarcoma
cells. J Mater Sci Mater Med 20, 347–50.
15. Tünnemann, G., Martin, R. M., Haupt, S.,
Patsch, C., Edenhofer, F., and Cardoso, M. C.
(2006) Cargo-dependent mode of uptake and
bioavailability of TAT-containing proteins and
peptides in living cells. FASEB J 20,
1775–84.
16. Duchardt, F., Fotin-Mleczek, M., Schwarz,
H., Fischer, R., and Brock, R. (2007) A com-
prehensive model for the cellular uptake of
cationic cell-penetrating peptides. Traffic 8,
848–66.
17. Palm-Apergi, C., Lorents, A., Padari, K.,
Pooga, M., and Hällbrink, M. (2009) The
membrane repair response masks membrane
disturbances caused by cell-penetrating pep-
tide uptake. FASEB J 23, 214–23.
18. Koppel, K., Padari, K., Lorents, A., Hällbrink,
M., Mano, M., Pedroso de Lima, M. C., and
Pooga, M. (2010) S413-PV cell-penetrating
peptide forms nanoparticle-like structures to
gain entry into cells. Bioconjug Chem 21,
774–83.
19. Hayat, M. A. Principles and techniques of
electron microscopy: biological applications.
Fourth edition: Cambridge University Press,
Cambridge; 2000.
20. Arai, R., Geffard, M., and Calas, A. (1992)
Intensification of labelings of the immuno-
gold silver staining method by gold toning.
Brain Res Bull 28, 343–5.