Synthesis and properties of differently charged chemiluminescent acridinium ester labels

Article abstract of DOI:10.1039/c2ob27190g

Chemiluminescent acridinium dimethylphenyl esters containing N-sulfopropyl groups in the acridinium ring are highly sensitive, hydrophilic labels that are used in automated immunoassays for clinical diagnostics. Light emission from these labels is triggered with alkaline peroxide in the presence of a cationic surfactant. At physiological pH, N-sulfopropyl acridinium esters exist as water adducts that are commonly referred to as pseudobases. Pseudobase formation, which results from addition of water to the zwitterionic N-sulfopropyl acridinium ring, neutralizes the positive charge on the acridinium nitrogen and imparts a net negative charge to the label due to the sulfonate moiety. As a consequence, N-sulfopropyl acridinium ester conjugates of small molecule haptens as well as large molecules such as proteins gain negative charges at neutral pH. In the current study, we describe the synthesis and properties of two new hydrophilic acridinium dimethylphenyl ester labels where the net charge in the labels was altered. In one label, the structure of the hydrophilic N-alkyl group attached to the acridinium ring was changed so that the pseudobase of the label contains no net charge. In the second acridinium ester, two additional negative charges in the form of sulfopropyl groups were added to the acridinium ring to make this label's pseudobase strongly anionic. Chemiluminescence measurements of these labels, as well as their conjugates of an antibody with a neutral pI, indicate that acridinium ester charge while having a modest effect on emission kinetics has little influence on light output. However, our results demonstrate that acridinium ester charge can affect protein pI, apparent chemiluminescence stability and non-specific binding of protein conjugates to microparticles. These results emphasize the need for careful consideration of acridinium ester charge in order to optimize reagent stability and performance in immunoassays. In the current study, we observed that for a neutral protein, an acridinium ester with a hydrophilic but charge-neutral N-alkyl group afforded faster light emission, lower non-specific binding and better chemiluminescence stability than an analogous label with an anionic N-alkyl group.

Full text of DOI:10.1039/c2ob27190g

Organic &
Biomolecular Chemistry
View Article Online
View Journal | View Issue
PAPER
Synthesis and properties of differently charged
chemiluminescent acridinium ester labels†
Cite this: Org. Biomol. Chem., 2013, 11,
1026
Anand Natrajan* and David Sharpe
Chemiluminescent acridinium dimethylphenyl esters containing N-sulfopropyl groups in the acridinium
ring are highly sensitive, hydrophilic labels that are used in automated immunoassays for clinical diagno-
stics. Light emission from these labels is triggered with alkaline peroxide in the presence of a cationic sur-
factant. At physiological pH, N-sulfopropyl acridinium esters exist as water adducts that are commonly
referred to as pseudobases. Pseudobase formation, which results from addition of water to the zwitterio-
nic N-sulfopropyl acridinium ring, neutralizes the positive charge on the acridinium nitrogen and imparts
a net negative charge to the label due to the sulfonate moiety. As a consequence, N-sulfopropyl acridi-
nium ester conjugates of small molecule haptens as well as large molecules such as proteins gain nega-
tive charges at neutral pH. In the current study, we describe the synthesis and properties of two new
hydrophilic acridinium dimethylphenyl ester labels where the net charge in the labels was altered. In one
label, the structure of the hydrophilic N-alkyl group attached to the acridinium ring was changed so that
the pseudobase of the label contains no net charge. In the second acridinium ester, two additional nega-
tive charges in the form of sulfopropyl groups were added to the acridinium ring to make this label’s
pseudobase strongly anionic. Chemiluminescence measurements of these labels, as well as their conju-
gates of an antibody with a neutral pI, indicate that acridinium ester charge while having a modest effect
on emission kinetics has little influence on light output. However, our results demonstrate that acridinium
ester charge can affect protein pI, apparent chemiluminescence stability and non-specific binding of
protein conjugates to microparticles. These results emphasize the need for careful consideration of acridi-
nium ester charge in order to optimize reagent stability and performance in immunoassays. In the
current study, we observed that for a neutral protein, an acridinium ester with a hydrophilic but charge-
neutral N-alkyl group afforded faster light emission, lower non-specific binding and better chemilumines-
cence stability than an analogous label with an anionic N-alkyl group.
Received 9th November 2012,
Accepted 10th December 2012
DOI: 10.1039/c2ob27190g
www.rsc.org/obc
completely understood, excited state acridone is believed to be
the light emitting species that is presumably formed from
Introduction
Chemiluminescent acridinium dimethylphenyl esters contain- addition of peroxide to C-9 of the acridinium ring (see Fig. 1,
ing N-sulfopropyl groups in the acridinium ring are highly sen- panel A for the numbering system) followed by scission of the
sitive, hydrophilic labels that are used in automated phenolic ester bond.2 Dioxetane and/or dioxetanone inter-
immunoassays for clinical diagnostics.1 Conjugates of these mediates have been proposed as immediate precursors of
acridinium esters are commonly used in conjunction with excited state acridone.2
magnetic microparticles in immunoassays for analytical
The surfactant CTAC influences light emission from acridi-
measurements of a wide range of clinically important analy- nium dimethylphenyl esters by two separate mechanisms.3 In
tes.1a,c,d Light emission from these labels and their conjugates the presence of CTAC, emission times from acridinium esters
is triggered with alkaline peroxide in the presence of the cat- and their conjugates are compressed from approximately 60
ionic surfactant cetyltrimethylammonium chloride (CTAC). seconds to ≤5 seconds for most labels and their conjugates.
While exact details of the light emission pathway are still not CTAC also enhances light yield by 3–4 fold by facilitating for-
mation of the dioxetane and/or dioxetanone intermediates.
These surfactant-mediated effects on light emission enable
Siemens Healthcare Diagnostics, Advanced Technology and Pre-Development, 333
fast assays with improved sensitivity and they require the
Coney Street, East Walpole, MA 02032, USA. E-mail: anand.natrajan@siemens.com;
binding of the acridinium ring to CTAC micelles through a
Fax: +1-508-660-4591; Tel: +1-508-660-4582
combination of hydrophobic and charge interactions.4
†Electronic supplementary information (ESI)
available. See DOI:
However, a label’s hydrophobicity and charge can exacerbate
10.1039/c2ob27190g
1026 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
Fig. 1 Panel A: Acridinium–pseudobase equilibrium of an N-sulfopropyl acridinium dimethylphenyl ester. Addition of water to C-9 neutralizes the positive charge
on the acridinium nitrogen and imparts a net negative charge to the label. Panel B: Structures of acridinium esters and their pseudobases of an anionic acridinium
ester 14 and charge-neutral and strongly anionic acridinium esters 9 and 13 respectively synthesized in the current study.
its non-specific binding to microparticles and can compromise
At physiological pH, acridinium esters with various N-alkyl
assay sensitivity in immunoassays. We recently reported4 that groups exist as water adducts that are commonly referred to as
the adverse effects of a hydrophobic acridinium ring can be pseudobases.4,5 Pseudobase formation, which results from the
mitigated by incorporating a highly polar, sulfobetaine zwitter- addition of water to the central ring of the acridinium ester,
ion linker in the phenolic ester. An example of such an opti- neutralizes the positive charge on the acridinium nitrogen.
mized structure is shown in Fig. 1 (compound 14, panel B). Consequently, triggering light emission from acridinium
This acridinium ester contains a relatively hydrophobic acridi- esters requires an acid pre-treatment step which restores the
nium ring with an electron-donating isopropoxy group at C-2 electrophilic center at C-9 by converting the pseudobase to the
and a phenolic ester with a sulfobetaine zwitterion linker. The acridinium form (Fig. 1, panel A). The pKa of acridinium to
isopropoxy group in 14 increases light output approximately pseudobase transition of acridinium esters depends upon the
twofold compared to the corresponding unsubstituted acridi- electronics of the acridinium ring and can be measured by
nium ester. Segregating the hydrophobic and hydrophilic spectrophotometry.4,5b For most acridinium esters, the pKa of
structural features in the acridinium ring and phenol respect- pseudobase formation is <7.4,6 Pseudobase formation in N-sulfo-
ively in 14 ensures low non-specific binding of the label to propyl acridinium esters (such as 14) imparts a net negative
microparticles without disrupting the critical influence of charge to the label due to the sulfonate moiety in the pseudo-
CTAC on its light emission.
base (Fig. 1, panel A). As a consequence, N-sulfopropyl
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1027

View Article Online
Paper
Organic & Biomolecular Chemistry
acridinium ester conjugates of small molecule haptens as well towards alkylating reagents10 with the exception of 1,3-
as large molecules such as proteins gain negative charges at propane sultone or strong methylating reagents, we decided to
pH ≥ 7.
use the easily-introduced N-sulfopropyl group (using our new
Despite the widespread use of N-sulfopropyl acridinium synthetic protocol9) as the scaffold to construct an acridinium
esters and analogous compounds7 in immunoassays, no study ester with a charge-neutral but highly polar N-alkyl group
has addressed the question of whether these negatively (compound 9, Fig. 1).
charged labels are really optimal in every situation? To answer
The synthetic scheme for compound 9 is illustrated in
this question, in the present study, we have investigated how Fig. 2. Compound 1, which is a synthetic intermediate of com-
overall charge of the acridinium ring impacts the properties of pound 144 with an N-sulfopropyl acridinium ring, was used as
chemiluminescent acridinium dimethylphenyl ester labels. We a starting point for the synthesis. Initially, we attempted to
describe the syntheses and properties of two new acridinium directly convert compound 1 to compound 5 by activating the
esters with differently charged (charge-neutral or strongly sulfonate moiety in 1 followed by condensation with the zwit-
anionic) acridinium rings in their respective pseudobases terionic amine derivative 4 whose synthesis we described
(Fig. 1, panel B, compounds 9 and 13). We also illustrate how recently.1d Caddick and coworkers11 have described a con-
these labels influence the properties of a monoclonal antibody venient method to convert sulfonate salts of heterocyclic
with a neutral pI in comparison to a conventional acridinium amines to sulfonamides and pentafluorophenyl sulfonate
ester label with an N-sulfopropyl acridinium ring (Fig. 1, panel esters using triphenylphosphine and trifluoromethanesulfonic
B, compound 14). Our results indicate that overall charge of anhydride for sulfonate activation. Treatment of compound 1
chemiluminescent acridinium ester labels can influence prop- with these reagents followed by pentafluorophenol however
erties such as pI, non-specific binding and apparent chemi- failed to give the pentafluorophenyl ester 3a. The poor solubi-
luminescence stability of protein conjugates. All are important lity of the zwitterionic amine 4 in the solvent (dichloro-
factors that can affect reagent performance in immunoassays.
methane) used for activation11 of the sulfonate
1 also
precluded any chance of reaction between 1 and 4. We next
resorted to the conventional method for activating sulfonates
which was to convert 1 to the sulfonyl chloride using neat
thionyl chloride. This activation reaction required careful
control of reaction time to prevent cleavage of the entire sulfo-
nate moiety and its replacement with chlorine. Short reaction
Results and discussion
Synthesis of acridinium ester labels and conjugates (Fig. 2 and
3, S1–S9,† Table 1)
At first glance, the synthesis of an acridinium ester whose times (≤30 minutes) minimized this side reaction although
pseudobase is charge-neutral appears trivial given that attempted condensation of the sulfonyl chloride with the zwit-
N-methyl acridinium phenyl esters and their derivatives were terionic amine 4 in dimethyl sulfoxide led mainly to hydrolysis
described more than 20 years ago.2a,b,8 The pseudobases of of the moisture sensitive sulfonyl chloride and no evidence of
these compounds do not contain any charge. However, the product 5.
acridinium ring in these compounds is very hydrophobic and
With these failed attempts to directly convert 1 to com-
overall aqueous solubility of these labels is quite low making pound 5, we decided to use a two step approach as illustrated
them generally unsuitable for labeling hydrophobic haptens.1a in Fig. 2. Condensation of the sulfonyl chloride of 1 with N,N-
When first described,1a acridinium dimethylphenyl esters with dimethylethylenediamine in acetonitrile led to the sulfona-
N-sulfopropyl groups in the acridinium ring represented an mide 2 which was isolated in 70% yield by preparative HPLC.
advance over N-methyl acridinium esters because they alle- The tertiary amine in 2 was then alkylated with 1,3-propane
viated the solubility problems of the latter and yet were sultone in the ionic liquid 1-butyl-3-methylimidazolium hexa-
sufficiently hydrophobic for efficient binding to CTAC micelles. fluorophosphate [BMIM][PF6] to give compound 5. This alkyl-
Also importantly, the N-sulfopropyl acridinium esters could be ation reaction in the ionic liquid gave complete conversion to
conveniently synthesized by N-alkylation of their acridine ester product 5 with no trace of starting material by HPLC analysis.
precursors with the potent alkylating reagent 1,3-propane sul- As an alternative to this two step approach, we also examined
tone.1a The main drawback of the original synthetic protocol conversion of the sulfonate of 1 to the more stable fluorophe-
was that the N-alkylation reaction had to be conducted at elev- nyl esters 3a and 3b and their subsequent coupling to the zwit-
ated temperatures with the toxic alkylating reagent serving the terionic amine 4. This approach appeared attractive for several
dual role of reactant and solvent. This was necessitated by the reasons. First, sulfonyl pentafluorophenyl esters are much
very poor reactivity of the acridine nitrogen of acridine more stable than sulfonyl chlorides and published studies
dimethylphenyl esters towards alkylating reagents. Recently,9 suggest that they can be coupled to amines even under
we reported that the N-alkylation of acridine dimethylphenyl aqueous conditions.12 Second, recent publications have
esters with and without various alkoxy groups in the acridine demonstrated that the reactivity of fluorophenyl esters of car-
ring with 1,3-propane sultone proceeds very efficiently in boxylic acids towards amine nucleophiles can be tuned by the
highly polar, aprotic, room temperature ionic liquids with sig- number of fluorines in the fluorophenyl ester.13 If the reacti-
nificantly reduced quantities of the alkylating reagent. Given vity of sulfonyl fluorophenyl esters can be tuned in a similar
the generally very poor reactivity of the acridine nitrogen manner, it can provide synthetic access to various
1028 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
Fig. 2 Synthetic scheme for compound 9. Reagents: (a) thionyl chloride; (b) N,N-dimethylethylenediamine, MeCN; (c) 1,3-propane sultone, 2,6-di-tert-butylpyri-
dine, 1-butyl-3-methylimidazolium hexafluorophosphate [BMIM][PF6]; (d) pentafluorophenol or 2,6-difluorophenol, triethylamine, dichloromethane; (e) diisopropyl-
ethylamine, DMSO; (f) 1 M HCl; (g) N,N,N’,N’-tetramethyl-O-(N-succinimidyl)uronium tetrafluoroborate (TSTU), diisopropylethylamine, DMF; (h) 0.25 M sodium
bicarbonate, DMF; (i) glutaric anhydride, diisopropylethylamine, DMF–water; ( j) TSTU, diisopropylethylamine, DMF–water.
N-propylsulfonamide-substituted acridinium esters. Finally, from
Synthesis of the fluorophenyl esters 3a and 3b was accom-
a practical point of view, we noticed that separation of starting plished via the sulfonyl chloride of 1 followed by condensation
material 1, fluorophenyl esters 3a and 3b, and product 5 by pre- with either pentafluorophenol or 2,6-difluorophenol. The
parative HPLC was much easier than separation of 5 from 2.
fluorophenyl esters 3a and 3b were isolated in 70–80% yield
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1029

View Article Online
Paper
Organic & Biomolecular Chemistry
Fig. 3 Synthetic scheme for compound 13. Reagents: (a) 1,3-propane sultone, sodium carbonate, [BMIM][PF6]; (b) 1 M HCl; (c) TSTU, diisopropylethylamine, DMF.
Table 1 Anti-HBsAg antibody labeling with acridinium esters 9, 13 and 14
Completion of the rest of the synthesis of acridinium ester
Input, equivalents
of acridinium ester
in labeling reaction
No. of labels
by mass
spectroscopy
Specific activity
(RLUs per
9 was relatively straightforward using chemistry we have
described previously.3,4 The methyl ester in 5 was cleaved by
acid hydrolysis and the carboxylic acid derivative 6 was isolated
in 72% yield by preparative HPLC. Condensation of 6 with the
zwitterionic diamine linker 73 was achieved using standard
N-hydroxysuccinimide ester chemistry for activation of the car-
boxylate 6. Compound 8, which was isolated in 85% yield by
preparative HPLC, was then condensed with glutaric anhydride
and the glutarate intermediate was subsequently converted to
Acridinium
ester label
mole × 10−19
)
9
9
13
13
14
14
5
10
5
10
5
3.0
5.0
3.0
6.0
3.0
6.0
6.0
6.4
6.3
6.1
6.2
4.2
10
after preparative HPLC and proved to be quite stable showing the target N-hydroxysuccinimide ester 9.
little signs of decomposition at room temperature. We next
Synthesis of the strongly anionic label 13 was achieved as
investigated reaction conditions for the coupling of these illustrated in Fig. 3. To preserve the strongly anionic nature of
fluorophenyl esters to the amine 4. While attempted coupling this compound, we decided not to introduce the neutral zwit-
of the pentafluorophenyl ester 3a to 4 under aqueous con- terionic linker 7 present in compounds 13 and 14 which have
ditions (0.25 M sodium bicarbonate–dimethyl sulfoxide) led more hydrophobic acridinium rings compared to 13. Pre-
mainly to hydrolysis of the pentafluorophenyl ester, coupling viously described 2,7-dihydroxyacridine ester derivative 1014
in neat dimethyl sulfoxide at 65–100 °C led to 60% conversion was alkylated in one step at both the two phenolic groups as
to product 5 with an isolated yield of 40% after preparative well as the acridine nitrogen with 1,3-propane sultone in the
HPLC. The 2′,6′-difluorophenyl ester 3b proved to be much ionic liquid [BMIM][PF6] with sodium carbonate or potassium
less reactive and showed no evidence of hydrolysis or coupling carbonate as the base. Conversion of 10 to the trisulfonated
to 4 under aqueous conditions at room temperature. However product 11 required the use of acid-free 1,3-propane sultone
in dimethyl sulfoxide at ∼100 °C, 70% conversion to product 5 and strict anhydrous conditions. The use of commercial 1,3-
was observed with an isolated yield of 50% by HPLC. Our propane sultone directly without distillation led to the for-
results suggest that for aliphatic sulfonates, 2,6-difluorofluoro- mation of mixtures of sulfonates that were difficult to separate.
phenyl esters may be superior to pentafluorophenyl esters due Synthesis of the target 13 was completed by acid hydrolysis of
to their increased stability especially in aqueous solvents. The the methyl ester in 11 which was isolated in 55% yield by pre-
synthetic routes that we have outlined should provide access to parative HPLC (two steps), followed by conversion of the car-
various N-propylsulfonamide-substituted acridinium com- boxylate 12 to the N-hydroxysuccinimide ester 13 using
pounds that would normally be very difficult to synthesize standard chemistry. The final product 13 was isolated in 93%
using alternate chemistry.
yield by preparative HPLC. HPLC analyses of all the
1030 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
intermediates and final compounds of Fig. 2 and 3 are shown
in the ESI (Fig. S1–S9†).
Protein conjugates of the labels 9, 13 and 14 were prepared
as described in the Experimental section using an in-house,
murine, anti-HBsAg monoclonal antibody with a neutral pI
(HBsAg = hepatitis B surface antigen). All three labels dis-
played similar reactivity towards the protein and the extent of
label incorporation was very similar as described in the Exper-
imental section (Table 1). Using an input of 5–10 equivalents
of the acridinium ester labels led to incorporation of 3–6
labels for each label as measured by MALDI-TOF (matrix-
assisted laser desorption ionization-time of flight) mass
spectroscopy.
Emission spectra, pH titrations and chemiluminescence
measurements (Fig. 4–7, S10†)
Fig. 5 pH titrations illustrating conversion of the acridinium form to the
pseudobase for compounds 9, 13 and 14. The bulky N-alkyl group in compound
9 shifted the pKa to acidic pH compared to compound 14. The strongly anionic
label 13 showed a shift in pKa to basic pH owing to increased electron donation
by the two ether oxygens at C-2 and C-7.
Emission spectra of acridinium ester labels 9, 13 and 14 are
shown in Fig. 4 along with the structures of the putative acri-
dones that are formed during the chemiluminescent reactions
of these labels. The emission spectra were recorded using a
spectral camera in a mostly aqueous medium (>90% water) as
described in the Experimental section. The emission spectrum
of 14 was reported in our previous study4 and is included here
for reference. The emission spectrum of the charge-neutral
label 9 was observed to be identical to that of 14 with an emis-
sion maximum centered at 458 nm. Both these acridinium
esters contain the same C-2 isopropoxy functional group in the
acridinium ring which is mainly responsible for the batho-
chromic shift in their emission spectra when compared to
unsubstituted N-sulfopropyl acridinium esters whose emission
maxima are typically observed at 430 nm.1c,4 The bulky, zwit-
terion-substituted, N-propylsulfonamide group in 9 was not
expected to affect its emission spectrum and Fig. 4 confirms
this expectation. On the other hand, the emission spectrum of
Fig. 6 Chemiluminescence emission profiles of compounds 9, 13 and 14 in the
presence of cetyltrimethylammonium chloride (CTAC). All three acridinium esters
showed rapid light emission in the presence of the cationic surfactant with
similar light yield (RLU = relative light unit).
compound 13 showed a bathochromic shift in the emission
spectrum with a maximum centered at 474 nm. This result is
consistent with our earlier observations on the chemilumines-
cence emission spectra of electron-rich, 2,7-dialkoxy-substi-
tuted acridinium esters that also showed bathochromic shifts
compared to monoalkoxy-substituted compounds.1c
Acridinium esters, at acidic pH, show strong long-wave-
length absorption bands due to the acridinium chromophore
that disappears as the pH is raised due to formation of the
pseudobase.4,5 Formation of the pseudobase by addition of
water at C-9 (Fig. 1, panel A) results in disruption in conju-
gation of the acridinium ring with concomitant loss of the
long wavelength absorption band. Pseudobase formation can
be followed by spectrophotometry and the pKa of the acridi-
nium to pseudobase transition can be determined by varying
the pH.4 The pKa of the acridinium to pseudobase transition
Fig. 4 Emission spectra of acridinium esters 9, 13 and 14. Structures of the
putative acridones are shown for the three acridinium esters. Due to similar C-2
alkoxy substitution, the emission spectrum of compound 9 was observed to be
identical to that of compound 14. Compound 13 showed a bathochromic shift
in its emission spectrum due to two electron-donating ether oxygens at C-2 and
C-7.
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1031

View Article Online
Paper
Organic & Biomolecular Chemistry
absence of CTAC and for 10 seconds integrated at 0.1 s intervals
in the presence of CTAC. The light collection time was
sufficiently long for complete emission under all conditions.
The emission profiles of the labels 9, 13 and 14 in the pres-
ence of CTAC and in its absence are shown in Fig. 6 and S10
(ESI†) respectively. Consistent with our earlier observations
with other acridinium ester labels,3,4 light emission in the
absence of CTAC was slow requiring more than a minute for
complete emission as illustrated in Fig. S10.† In the presence
of CTAC, these emission times were compressed to ≤5 seconds
(Fig. 6). These results indicate that the acridinium rings of the
three labels partition strongly into CTAC micelles.4 Despite
differences in their pKa for pseudobase formation, in the pres-
ence of excess peroxide ions, emission kinetics of the three
labels were quite similar. Specific activity of the labels in the
presence of CTAC was also observed to be quite similar with
observed specific activities of 5.9, 5.4 and 4.6 × 1019 RLUs per
mole (RLU = relative light unit) for 9, 13 and 14 respectively
(average of three replicates). Total light yield was increased
approximately 3-fold for each label in the presence of CTAC.3,4
Chemiluminescence emission profiles of the protein conju-
gates are shown in Fig. 7. Light emission was again observed
to be rapid for all three labels and was not affected by label
density. Modest differences in the emission kinetics were
noted for the three labels with the conjugates of 9 showing the
fastest light emission (>90% emission in 0.5 second) whereas
conjugates of the strongly anionic label 13 were the slowest
emitters (>90% emission in 2 seconds). Emission kinetics of
the conjugates of label 14 fell in between requiring one second
for >90% emission. Despite these differences in emission kin-
etics, the specific activities of the labels in the conjugates were
observed to be quite similar as illustrated in Table 1 (Experi-
mental section).
In an earlier empirical study, we reported that acridinium
esters with methoxy groups at C-2 and/or C-7 display the great-
est increase in light yield (2–3-fold) compared to other regioi-
someric, methoxy-substituted acridinium esters.1c It has been
suggested in recent publications that competition between
hydroperoxide ions and hydroxide ions for reaction at C-9 of
acridinium esters drives the light and dark reactions respecti-
vely2d and that these dark reactions are suppressed for 2,7-
dimethoxy-substituted acridinium esters.15 The pKa of acridi-
nium to pseudobase transition is shifted progressively to more
basic pH for these alkoxy-substituted esters as observed in this
study and our earlier reports.4 Intuitively, an increase in pKa
should reduce the electrophilic character at C-9 and increase
discrimination between more nucleophilic hydroperoxide ions
(light reaction) rather than hydroxide ions (dark reaction).
However, as observed for compound 9, a decrease in pKa is not
always correlated with a drop in light yield. Similarly, an
increase in pKa for compound 13 did not increase light output.
Decomposition of dioxetane intermediates formed in the
chemiluminescent reaction of acridinium esters is postulated
to occur via electron transfer from the acridine nitrogen2b,16 in
a mechanism that is analogous to Schuster’s CIEEL (chemi-
cally-initiated electron exchange luminescence) mechanism.17
Fig. 7 Chemiluminescence emission profiles of anti-HBsAg antibody conju-
gates of acridinium esters 9, 13 and 14 in the presence of CTAC. Label density
was observed not to affect emission kinetics or light yield for each label. Emis-
sion kinetics of the conjugates increased in the order 9 > 14 > 13 (HBsAg =
hepatitis B surface antigen).
of acridinium esters typically reflects the electron density at
C-9 of the acridinium ring. Electron donating groups at C-2
and/or C-7 decrease the electron density at C-9 and shift the
pKa for pseudobase formation to basic pH.4,6 pH titrations of
the acridinium esters 9, 13 and 14 are shown in Fig. 5. As
expected, the 2,7-dialkoxy-substituted acridinium ester 13 dis-
played a shift in the pKa for pseudobase formation to 6.5 com-
pared to 14 (pKa ∼ 4.5) which has a single C-2 alkoxy group.
Surprisingly, acridinium ester 9, despite being substituted
with the same C-2 isopropoxy group as 14, exhibited a shift in
the pKa to acidic pH (pKa ∼ 4). These observations suggest that
the bulky N-alkyl group in 9 disfavors a flat acridinium ring
due to steric congestion. Formation of the pseudobase pre-
sumably relieves this steric congestion. We have observed that
other acridinium esters with bulky N-alkyl groups also exhibit
shifts in their pKa for pseudobase formation to acidic pH
(unpublished data). Our observations also imply that the pKa
of pseudobase formation of acridinium esters is indicative of
the electron density at C-9 only when the N-alkyl groups in the
acridinium ring are identical.
Chemiluminescence measurements of the labels 9, 13 and
14 as well as their anti-HBsAg antibody conjugates were
carried out as described previously.3,4 In a typical experiment,
1–2 mg mL−1 solutions of HPLC-purified labels and the corre-
sponding protein conjugates of these compounds were serially
diluted in phosphate buffer to approximately 0.5 nM (nM =
nanomolar, 10−9 M) for the free labels, and approximately
0.2 nM for the protein conjugates as described in the Experi-
mental section. Chemiluminescence from 0.01 mL of each
diluted sample was triggered by the addition of 0.3 mL each of
0.1 M nitric acid containing 0.5% hydrogen peroxide followed
by the addition of 0.25 M sodium hydroxide. In experiments
with CTAC, the surfactant was included in the second reagent
(0.25 M sodium hydroxide) at five times its reported critical
micelle concentration (CMC) in water.3,4 Light was collected for
a total of two minutes, integrated at 0.5 second intervals in the
1032 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
This electron transfer mechanism may be more efficient for
electron rich C-2 and/or C-7 alkoxy-substituted acridinium
esters and may explain why compounds 9, 13 and 14 have the
same light output despite the observed differences in their pKa
for pseudobase formation.
Conjugate pI, chemiluminescence stability and non-specific
binding (Fig. 8, 9, S11,† Table 2)
The acridinium rings of the labels 9, 13 and 14 have different
charge characteristics in their respective pseudobases and we
examined their impact on protein pI, chemiluminescence stab-
ility as well as non-specific binding to magnetic microparticles
that are commonly used in immunoassays.
Fig. 9 Apparent chemiluminescence stability of anti-HBsAg antibody conju-
gates of labels 9, 13 and 14 at pH 6.0 and pH 7.4 at 37 °C. Order of stability at
pH 7.4 was observed to be 9 > 14 > 13 which was inversely correlated with the
pKa for pseudobase formation. Loss of chemiluminescence was not observed at
either pH for all three labels at 4 °C (Fig. S11†).
To study the effect of these differently charged labels on
protein pI, we selected a monoclonal antibody with a neutral
pI (an in-house anti-HBsAg antibody) so that small pertur-
bations in the charge of the protein caused by acridinium
ester labeling could be more easily measured. Isoelectric focus-
ing (IEF) gel electrophoresis is a commonly-used technique18
to measure protein pI and we used this method to study the
impact of the three labels 9, 13 and 14 on the pI of the anti-
HBsAg antibody conjugates (Fig. 8). As shown in Fig. 8, label-
ing with the anionic label 14 and the strongly anionic label 13
shifted the pI of the protein from approximately 7 (lane 3) to
approximately 5.5 for both labels (lanes 4–7). An increase in
label density of either of these two labels showed more
compact bands in the gel and a slightly greater shift in protein
pI to acidic pH. Interestingly, although label 13 contains two
additional negative charges compared to 14, the IEF gel was
unable to resolve any differences in pI in the conjugates of
these two labels. Nevertheless, these results nicely illustrate
the effect of these negatively charged labels on altering the pI
of a neutral protein. In contrast, conjugates of the charge-
neutral label 9 showed only a small shift in the pI of the
protein to acidic pH (Fig. 8, lanes 8 and 9). The small shift is
attributable to neutralization of some of the lysine residues on
the protein caused by acridinium ester labeling. Thus, as pre-
dicted by its structural features, acridinium ester 9 contains no
net charge in the acridinium ring and therefore had a minimal
effect on protein pI.
Non-specific binding is a better reflection of molecular
forces and we measured the non-specific binding of the more
heavily-labeled conjugates (Tables 1 and 2) of 9, 13 and 14 to
three different types of magnetic particles that are commonly
used in immunoassays. The particles selected were paramag-
netic particles (PMPs) and two commercially available mag-
netic latex particles (MLPs). The PMPs used were 1–10 micron-
sized iron(III) oxide particles, coated with a murine anti-fluo-
rescein antibody which was coupled to the amino-silanized
particle surface using glutaraldehyde. The two MLPs (Dyna-
beads® M-270 and M-280 particles) are available from a com-
mercial vendor (Life Technologies). Both particles were
2.8 micron in size and were coated with streptavidin. The
intrinsic, surface characteristics of the three types of particles
are quite different because of different functional groups.
PMPs contain amines on their surfaces whereas M-270 par-
ticles have hydrophilic carboxylate surfaces. The M-280 par-
ticles have a more hydrophobic surface with a tosylate-
activated polystyrene coating (Experimental section). The two
types of particles (PMPs and MLPs) were mixed with solutions
Fig. 8 Isoelectric focusing (IEF) gel of anti-HBsAg antibody conjugates with
varying label density of acridinium esters 9, 13 and 14. The pI of the neutral
antibody was shifted to acidic pH by the anionic labels 13 and 14 whereas the of the acridinium ester conjugates and were then magnetically
charge-neutral label 9 had a much smaller effect on protein pI. Lanes: (1) Novex
separated. They were subsequently washed twice, either with
water or a surfactant solution, and then the chemilumines-
cence associated with the particles was measured. The ratio of
IEF markers pH 3–10; (2) blank; (3) unlabeled antibody; (4) antibody conjugate
of 13, 3 labels; (5) antibody conjugate of 13, 6 labels; (6) antibody conjugate of
14, 3 labels; (7) antibody conjugate of 14, 6.0 labels; (8) antibody conjugate of
this chemiluminescence value in comparison to the total che-
miluminescence input is referred to fractional non-specific
9, 3 labels; (9) antibody conjugate of 9, 5 labels; (10) Novex IEF markers pH
3–10.
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1033

View Article Online
Paper
Organic & Biomolecular Chemistry
Table 2 Fractional non-specific binding (FNSB) of acridinium ester (9, 13 and 14) conjugates of anti-HBsAg antibody conjugates to three different magnetic micro-
particles measured on a Siemens Healthcare Diagnostics’ ADVIA Centaur® system
Particles
14, surfactant wash
14, water wash
9, surfactant wash
9, water wash
13, surfactant wash
13, water wash
PMP
M-280-SA
M-270-SA
2.6 × 10−4
1.3 × 10−4
1.8 × 10−5
2.4 × 10−4
1.3 × 10−4
1.8 × 10−5
2.0 × 10−4
6.6 × 10−5
1.6 × 10−5
2.1 × 10−4
6.5 × 10−5
1.6 × 10−5
1.5 × 10−4
4.9 × 10−5
1.3 × 10−5
1.4 × 10−4
4.8 × 10−5
0.9 × 10−5
An input of 250–300 million RLUs, 5–6 labels per antibody corresponding to 1 picomole (1 picomole = 10−12 mole) of anti-HBsAg antibody
conjugate and 70 μg of magnetic particles was used for each measurement (ten replicates). Surfactant wash corresponded to 0.8 mM Triton X-100
in PBS buffer, pH = 7.2. Abbreviations used: RLU = relative light unit; PMP = paramagnetic particle; M-280 and M-270 are commercial names of
Dynabeads® (Life Technologies) with hydrophobic, tosyl-activated and hydrophilic carboxylate surfaces respectively; SA = streptavidin.
binding (FNSB) and reflects the resistance of the conjugate of increasing pKa for pseudobase formation for these labels. At
towards non-specific adsorption to the microparticles. The first, these observations appear counterintuitive because an
results of these measurements are tabulated in Table 2 and increase in pKa should decrease the positive charge at C-9 and
details are described in the Experimental section.
stabilize the adjacent phenolic ester linkage. However, at a
The results in Table 2 indicate that the FNSB values of the given pH, an increase in pKa for pseudobase formation causes
anti-HBsAg antibody conjugates of the labels 9, 13 and 14 are a greater proportion of the acridinium ester to be in the acridi-
not only dependent upon the charge of the acridinium ester nium form which is expected to be more susceptible to
label, but are also dependent upon the surface characteristics hydrolysis of the phenolic ester due to electron withdrawal
of the particles used in the measurements. For all three par- from C-9. At pH 6, the charge-neutral label 9 with the lowest
ticles, regardless of the wash method (water or surfactant sol- pKa was still the most stable. But the order of stability of 13
ution wash), FNSB of the labels decreased in the order 13 and 14 was reversed with 13 (higher pKa) being more stable
(strongly anionic) < 9 (charge-neutral) < 14 (anionic) defying than 14. The apparent anomalous behavior of the strongly
conventional wisdom that non-ionic but hydrophilic func- anionic label 13 may be the result of a combination of several
tional groups are best for preventing protein adsorption to sur- factors. The label is expected to be strongly anionic (because of
faces.19 The order of non-specific binding on the three types of the two additional sulfonates in the acridinium ring at C-2 and
particles was observed to be M-270 < M-280 < PMP for all three C-7) at both pH = 6 and 7.4 so should repel negatively charged
labels. Our results (Table 2) clearly demonstrate that acridi- ions such as hydroxide ions more effectively than 14. Yet, the
nium ester charge does play an important role in dictating an acridinium ester also contains an electrophilic center at C-9
important property of immunoassay reagents, i.e. non-specific (in the acridinium form) that can destabilize the adjacent phe-
binding. The strongly anionic label 13 appears to be the best nolic ester. At pH 6, a greater proportion of this acridinium
label for a neutral protein based on these measurements but ester is in the labile acridinium form compared to 14 (Fig. 5)
the addition of multiple negative charges may not be condu- but the concentration of hydroxide ions is low. At pH 7.4, the
cive to assay kinetics that use acidic proteins and particles label is still strongly anionic but there is less labile acridinium
with negatively charged surfaces such as the M-270 particles. form despite a greater concentration of hydroxide ions. These
Acridinium ester 9 may be the more attractive label for all opposing factors may be responsible for the unusual pH-
three types of particles because it is charge-neutral yet hydro- dependent stability behavior of 13 with the lower concen-
philic with lower FNSB compared to the conventional anionic tration of hydroxide ions and charge repulsion dictating better
label 14. This is most evident for the hydrophobic M-280 par- stability at pH = 6.0 despite a higher proportion of the labile
ticles where the FNSB of 9 was twofold lower than 14. This acridinium form at this pH.
observation indicates that the acridine ring in the pseudobase
of the charge-neutral label 9 is more hydrophilic than that of
Conclusions
the anionic label 14. Moreover, because of its unique structural
features, as illustrated here with the neutral anti-HBsAg anti-
body, acridinium ester 9 is also not expected to alter the
charge characteristics of proteins or small haptens.
Finally, we address acridinium ester chemiluminescence
stability which is another important property of these labels
for immunoassay applications. Acridinium ester instability is
mostly attributed to hydrolysis of the phenolic ester linkage.8
Apparent chemiluminescence stability of anti-HBsAg antibody
conjugates of the three labels 9, 13 and 14 at pH = 6 and 7.4 at
37 °C is shown in Fig. 9. (No differences in stability could be
discerned at 4 °C as shown in Fig. S11†) At pH 7.4, the order of
stability was observed to be 9 > 14 > 13 which is also the order
We have described the syntheses of two new, differently
charged chemiluminescent acridinium dimethylphenyl esters
9 and 14 using synthetic routes that we developed specifically
for assembling these unusual structures. Chemiluminescence
measurements of these labels as well as their conjugates of a
protein with a neutral pI indicated modest differences in emis-
sion kinetics but similar light yield. These differently charged
labels however showed significant differences in both non-
specific binding to microparticles and how they impacted
protein pI. The labels also exhibited significant pH-dependent
differences in chemiluminescence stability. Although not a
focus of the current study, these differently charged acridi-
nium ester labels may also influence other immunoassay
1034 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
parameters such as binding rates and affinity constants of con- nitrogen atmosphere for 30 minutes. The reaction was then
jugates which will need to be evaluated separately for each cooled to room temperature, diluted with anhydrous toluene
assay. Our results emphasize the need for careful consider- (5 mL) and concentrated under reduced pressure. The crude
ation of acridinium ester charge in order to optimize reagent sulfonyl chloride was then treated with an ice-cold solution of
stability and performance in immunoassays that use these che- N,N-dimethylethylenediamine (0.45 mL, 4.1 mmoles) in anhy-
miluminescent labels. Because a multitude of factors can drous MeCN (5 mL). The resulting solution was stirred at 0 °C
affect immunoassay performance, empirical approaches such under a nitrogen atmosphere. After 15 minutes, the reaction
as the one we have described have considerable merit. During was analyzed by HPLC using a Phenomenex, C18, 10 micron,
the course of this study, we have identified a new structural 3.9 × 300 mm column and a 30 minute gradient of 10 → 100%
motif for acridinium esters, i.e. a charge-neutral, zwitterion- MeCN–water (each with 0.05% TFA, TFA = trifluoroacetic acid)
substituted hydrophilic acridinium ring, such as that in acridi- at a flow rate of 1.0 mL per minute and UV detection at
nium ester 13. This new acridinium ester 13, when conjugated 260 nm. Product was observed eluting at 17 minutes with no
to an antibody with a neutral pI showed faster emission kin- starting material at 19 minutes. The reaction was diluted with
etics, lower non-specific binding and better chemilumines- toluene (5 mL) and concentrated under reduced pressure. The
cence stability when compared to the analogous label 14 with crude reaction mixture was dissolved in DMF (4 mL) and puri-
an N-sulfopropyl acridinium ring.
fied by preparative HPLC using a YMC, 10 micron, C18, 30 ×
250 mm column and a 30 minute gradient of 10 → 100%
MeCN–water (each with 0.05% TFA) at a solvent flow rate of
20 mL per minute and UV detection at 260 nm. The HPLC frac-
tions containing product 2 were combined and concentrated
under reduced pressure to yield a dark brown, sticky solid.
Yield = 85 mg (70%). νmax/cm−1 3424 (NH), 1752 and 1680
(CO), 1471, 1326, 1202, 1144. δH (500 MHz, CF3COOD) 1.51 (d,
6H, J = 6.0), 2.54 (s, 6H), 2.88 (br s, 2H), 3.14 (s, 6H), 3.51 (br s,
2H), 3.80 (m, 4H), 4.07 (s, 3H), 4.91 (spt, 1H, J = 6.0), 5.73 (m,
2H) , 7.87 (br s, 1H), 8.00 (s, 2H), 8.09 (m, 1H), 8.20 (m, 1H),
8.43 (m, 1H), 8.66 (m, 2H), 8.72 (d, 1H, J = 9.0). MALDI-TOF
MS m/z 636.9 M+; HRMS m/z 636.2752 M+ (636.2743
calculated).
Compound 3a. Compound 1 (60 mg, 0.11 mmole) in thionyl
chloride (2 mL) was heated at 85 °C in an oil bath under a
nitrogen atmosphere for 30 minutes. The reaction was then
cooled to room temperature and hexanes (30 mL) were added
to precipitate the acid chloride. The hexanes were decanted
and the residue was rinsed with additional hexanes (2 ×
10 mL). The crude sulfonyl chloride was then treated with an
ice-cold solution of pentafluorophenol (94 mg, 0.51 mmole)
and triethylamine (0.072 mL, 0.52 mmole) in anhydrous
dichloromethane (5 mL). The resulting solution was stirred at
0 °C under a nitrogen atmosphere. After 1 hour, the reaction
was analyzed by HPLC using a Phenomenex, C18, 10 micron,
3.9 × 300 mm column and a 30 minute gradient of 10 → 100%
MeCN–water (each with 0.05% TFA) at a flow rate of 1.0 mL
per minute and UV detection at 260 nm. Compound 3a was
observed eluting at 25 minutes. The reaction was then concen-
trated under reduced pressure and the crude product was dis-
solved in MeCN (5 mL). The product was isolated by
preparative HPLC as described for compound 2. Yield = 55 mg
(71%); red hygroscopic solid. νmax/cm−1 1719 and 1688 (CO),
1522, 1474, 1367, 1325, 1186, 1147. δH (500 MHz, CF3COOD)
1.47 (d, 6H, J = 6.0), 2.50 (s, 6H), 2.99 (br s, 2H), 4.02 (s, 3H),
4.07 (br s, 2H), 4.87 (spt, 1H, J = 6.0), 5.76 (br s, 2H), 7.83 (s,
1H), 7.95 (s, 2H), 8.06 (m, 1H), 8.17 (br d, 1H), 8.41 (m, 1H),
8.68 (m, 3H); δF (470 MHz, CF3COOD) −154.3 (d, 2F, J = 16.9),
Experimental
General
Chemicals were purchased from Sigma-Aldrich (Milwaukee, Wis-
consin, USA) unless indicated otherwise. The ionic liquid [BMIM]
[PF6] was dried under high vacuum over P2O5 prior to use.
All final acridinium esters and intermediates were analyzed
and/or purified by HPLC using a Beckman-Coulter HPLC
system. Flash chromatography was performed using an ‘Auto-
flash’ system from ISCO. MALDI-TOF (matrix-assisted laser
desorption ionization-time of flight) mass spectroscopy was
performed using a Voyager DE™ Biospectrometry™ Worksta-
tion from Perkin-Elmer. This is a benchtop instrument operat-
ing in the linear mode with a 1.2 meter ion path length, flight
tube. Spectra were acquired in the positive ion mode. For
small molecules, α-cyano-4-hydroxycinnamic acid was used as
the matrix and spectra were acquired with an accelerating
voltage of 20 000 volts and a delay time of 100 ns. For protein
conjugates, sinapinic acid was used as the matrix and spectra
were acquired with an accelerating voltage of 25 000 volts and
a delay time of 85 ns.
For HRMS (high resolution mass spectra), samples were dis-
solved in HPLC-grade methanol and analyzed by direct-flow
injection (injection volume = 5 μL) electrospray ionization (ESI)
on a Waters Qtof API US instrument in the positive ion mode.
Optimized conditions were as follows: capillary = 3000 kV,
cone = 35, source temperature = 120 °C, desolvation tempera-
ture = 350 °C. NMR spectra were recorded on a Varian
500 MHz spectrometer. IR spectra of neat samples were
recorded on a Bruker TENSOR37 FT-IR spectrometer, ATR
mode on a ZnSe crystal. UV-Visible spectra were recorded on a
Beckman DU 7500 spectrophotometer. Chemiluminescence
measurements were carried out using a Berthold Technologies’
AutoLumat Plus LB953 luminometer.
1. Synthesis of acridinium esters (Fig. 2, 3 and Fig. S1–S9 ESI†)
Compound 2. Compound 1 (108 mg, 0.19 mmole) in thionyl −156.2 (t, 2F, J = 19.4), −163.0 (br t, 2F); MALDI-TOF MS m/z
chloride (4 mL) was heated at 85 °C in an oil bath under a 732.7 M+; HRMS m/z 732.1707 M+ (732.1691 calculated).
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1035

View Article Online
Paper
Organic & Biomolecular Chemistry
Compound 3b was prepared similarly. Yield = 58 mg (81%); to a 100 mL round bottom flask with methanol and concen-
red hygroscopic solid. νmax/cm−1 1710 and 1687 (CO), 1480, trated under reduced pressure. The residue was washed with
1377, 1198, 1164, 1145. δH (500 MHz, CF3COOD) 1.47 (d, 6H, toluene (3 × 20 mL) and then dried under vacuum. Crude
J = 6.0), 2.50 (s, 6H), 3.00 (br s, 2H), 4.03 (s, 3H), 4.05 (br s, product 5 was used directly for the next reaction.
2H), 4.87 (spt, 1H, J = 6.0), 5.76 (br s, 2H), 7.04 (br t, 2H), 7.31
Compound 6. The crude product 5 from above was sus-
(m, 1H), 7.83 (s, 1H), 7.95 (s, 2H), 8.06 (br t, 1H), 8.17 (br d, pended in 1 M HCl (10 mL) and was refluxed under a nitrogen
1H), 8.40 (m, 1H), 8.60 (br d, 3H). δF (470 MHz, CF3COOD) atmosphere. After 2 hours, the reaction was cooled to room
−127.5 (br t, 2F). MALDI-TOF MS m/z 678.6; M+; HRMS m/z temperature and analyzed by HPLC using a Phenomenex, C18,
678.1970 M+ (678.1973 calculated).
Compound 5
10 micron, 3.9 × 300 mm column and a 30 minute gradient of
10 → 70% MeCN–water (each with 0.05% TFA) at a flow rate of
1.0 mL per minute and UV detection at 260 nm. Clean conver-
sion to product 6 eluting at 19 minutes was observed. The
product was purified by preparative HPLC as described for
(a) Synthesis from compound 3a. A mixture of compound 3a
(40 mg, 0.055 mmole), compound 4 (29 mg, 0.138 mmole) and
diisopropylethylamine (0.024 mL, 0.138 mmole) in dimethyl
sulfoxide (2 mL) was heated at 65 °C under a nitrogen atmos-
phere for 2 hours. The reaction was then cooled to room temp-
erature and analyzed by HPLC using a Phenomenex, C18,
10 micron, 3.9 × 300 mm column and a 30 minute gradient of
10 → 100% MeCN–water (each with 0.05% TFA) at a flow rate
of 1.0 mL per minute and UV detection at 260 nm. Product 5
was observed eluting at 17 minutes (60% conversion) with
hydrolyzed starting material 1 (∼40%) eluting at 19 minutes.
The product 5 was purified by preparative HPLC as described
for compound 2. Yield = 17 mg (41%); red powder. νmax/cm−1
3443 (NH), 1720 and 1688 (CO), 1470, 1326, 1200, 1147. δH
(500 MHz, CF3COOD) 1.48 (d, 6H, J = 6.0), 2.52 (br s, 8H), 2.85
(m, 2H), 3.27 (s, 6H), 3.39 (br t, 2H), 3.67–3.96 (m, 8H), 4.05 (s,
3H), 4.88 (spt, 1H, J = 6.0), 5.70 (br s, 2H), 7.84 (s, 1H), 7.97 (s,
2H), 8.06 (m, 1H), 8.18 (br d, 1H), 8.41 (m, 1H), 8.64 (m, 2H),
8.69 (d, 1H, J = 8.5). MALDI-TOF MS m/z 758.2 M+; HRMS m/z
758.2753 M+ (758.2781 calculated).
(b) Synthesis from compound 3b. A mixture of compound
3b (30 mg, 0.044 mmole), compound 4 (46 mg, 0.22 mmole)
and diisopropylethylamine (0.078 mL, 0.44 mmole) in
dimethyl sulfoxide (3 mL) was heated at 95–100 °C under a
nitrogen atmosphere for 16 hours. The reaction was then
cooled to room temperature and analyzed by HPLC using a
Phenomenex, C18, 10 micron, 3.9 × 300 mm column and a
30 minute gradient of 10 → 100% MeCN–water (each with
0.05% TFA) at a flow rate of 1.0 mL per minute and UV detec-
tion at 260 nm. Product 5 was observed eluting at 17 minutes
(70% conversion) with hydrolyzed starting material eluting 1 at
19 minutes (16%) and unreacted 3b eluting at 24 minutes
(13%). The product 5 was purified by preparative HPLC as
described for compound 2. Yield = 15 mg (50%); red hygro-
scopic solid.
compound 2. Yield = 72 mg (72%); red hygroscopic solid. νmax
/
cm−1 3435 (OH and NH), 1749 and 1667 (CO), 1483, 1308,
1200, 1152. δH (500 MHz, CF3COOD) 1.51 (d, 6H, J = 6.0), 2.56
(br s, 8H), 2.87 (br s, 2H), 3.29 (s, 6H), 3.42 (br t, 2H),
3.69–4.02 (m, 8H), 4.92 (spt, 1H, J = 6.0), 5.74 (m, 2H), 7.87 (br
s, 1H), 8.07 (s, 2H), 8.09 (m, 1H), 8.22 (m, 1H), 8.44 (m, 1H),
8.67 (m, 2H), 8.72 (d, 1H, J = 8.0). MALDI-TOF MS m/z 745.5
(M + H)+, HRMS m/z 744.2644 M+ (744.2625 calculated).
Compound 8. A solution of compound
6
(72 mg,
0.097 mmole) in DMF (2 mL) and water (0.1 mL) was treated
with diisopropylethylamine (0.034 mL, 0.2 mmole) and N,N,N′,N′-
tetramethyl-O-(N-succinimidyl)uronium
tetrafluoroborate
(TSTU) (58 mg, 0.19 mmole). The reaction was stirred at room
temperature. After 15 minutes, the reaction was analyzed by
HPLC using a Phenomenex, C18, 10 micron, 3.9 × 300 mm
column and a 30 minute gradient of 10 → 70% MeCN–water
(each with 0.05% TFA) at a flow rate of 1.0 mL per minute and
UV detection at 260 nm. Clean conversion to the N-hydroxysuc-
cinimide (NHS) ester of 6 eluting at 21.3 minutes was
observed. The solution of the NHS ester was cooled to 4 °C
and was added dropwise to an ice-cold solution of compound
73 (0.216 g, 0.48 mmole, HBr salt) dissolved in a mixture
(5 mL) of 0.25 M sodium bicarbonate (85%) and DMF (15%).
The reaction was warmed to room temperature and stirred for
30 minutes. HPLC analysis of the reaction mixture as
described above showed clean conversion to product 8 eluting
at 14 minutes. The product was purified by preparative HPLC
as described for compound 2. Yield = 82 mg (85%); red hygro-
scopic solid. νmax/cm−1 3446 (NH), 1748 and 1662 (CO), 1473,
1202, 1176. δH (500 MHz, CF3COOD) 1.52 (d, 6H, J = 6.0), 2.39
(br m, 2H), 2.55 (br s, 12H), 2.88 (br s, 2H), 3.23 (s, 3H), 3.31
(s, 6H), 3.43 (br m, 6H), 3.55–4.06 (m, 16H), 4.91 (spt, 1H, J =
(c) Synthesis from compound 2. A mixture of compound 2 6.0), 5.74 (m, 2H), 7.70 (br s, 2H), 7.86 (br s, 1H), 8.10 (m, 1H),
(85 mg, 0.134 mmole), distilled 1,3-propane sultone (0.25 g, 8.22 (m, 1H), 8.45 (m, 1H), 8.67 (m, 2H), 8.72 (d, 1H, J = 9.0);
2.05 mmoles) and 2,6-di-tert-butylpyridine (0.22 mL, MALDI-TOF MS m/z 994.5 (M + H)+, HRMS m/z 993.4141 M+
1.0 mmole) in [BMIM][PF6] (1.0 mL) was heated at 150–155 °C (993.4136 calculated).
in a sealed tube for 16 hours. The reaction was then cooled to
Compound 9. A solution of compound 8 (62 mg,
room temperature and analyzed by HPLC using a Pheno- 0.062 mmole) in DMF (3 mL) and water (0.3 mL) was treated
menex, C18, 10 micron, 3.9 × 300 mm column and a 30 minute with diisopropylethylamine (0.054 mL, 0.31 mmole) and gluta-
gradient of 10 → 70% MeCN–water (each with 0.05% TFA) at a ric anhydride (36 mg, 0.31 mmole). The reaction was stirred at
flow rate of 1.0 mL per minute and UV detection at 260 nm. room temperature. After 30 minutes, the reaction was analyzed
Product 5 was observed eluting at 23 minutes with no trace of by HPLC using a Phenomenex, C18, 10 micron, 3.9 × 300 mm
starting material 2 at 22 minutes. The reaction was transferred column and a 30 minute gradient of 10 → 70% MeCN–water
1036 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
(each with 0.05% TFA) at a flow rate of 1.0 mL per minute and 8.75 (d, 2H, J = 10.0). MALDI-TOF MS m/z 769.4 M+; HRMS m/z
UV detection at 260 nm. The glutarate derivative of compound 770.1255 (M + H)+ (770.1247 calculated).
9 was observed eluting at 15.3 minutes (>80% conversion). The
Compound 13. A solution of compound 12 (10 mg,
reaction mixture was then treated with diisopropylethylamine 0.013 mmole) in DMF (1.5 mL) was treated with diisopropyl-
(0.13 mL, 0.75 mmole) and TSTU (0.26 g, 0.86 mmole). The ethylamine (0.0068 mL, 0.04 mmole) and TSTU (6 mg,
reaction was stirred at room temperature. After 30 minutes, 0.02 mmole). The reaction was stirred at room temperature.
the reaction was analyzed by HPLC as described above. After 30 minutes, the reaction was analyzed by HPLC using a
Product 9 was observed eluting at 16.5 minutes (>80% conver- Phenomenex, C18, 10 micron, 3.9 × 300 mm column and a
sion). The product was purified by preparative HPLC as 30 minute gradient of 10 → 70% MeCN–water (each with
described for compound 2. The HPLC fractions containing 0.05% TFA) at a flow rate of 1.0 mL per minute and UV detec-
product 9 were frozen at −80 °C and lyophilized. Yield = 42 mg tion at 260 nm. Product 13 was observed eluting at
(56%); fluffy reddish-yellow solid. νmax/cm−1 1737, 1705, 1688 14.5 minutes. The product was purified by preparative HPLC
and 1680 (CO), 1202, 1136. δH (500 MHz, CF3COOD) 1.55 (d, as described for compound 2. The HPLC fractions containing
6H, J = 6.0), 2.06–2.47 (m, 4H), 2.58 (br s, 10H), 2.67 (t, 2H, J = product were frozen at −80 °C and lyophilized. Yield = 8.3 mg
7.0), 2.83 (t, 2H, J = 7.0), 2.86–2.96 (m, 2H), 3.08 (s, 4H), 3.22 (93%); dark yellow solid. νmax/cm−1 3443 (OH), 1762, 1738 and
(s, 3H), 3.33 (s, 6H), 3.45 (m, 4H), 3.59 (m, 6H), 3.71–3.90 (m, 1615 (CO), 1464, 1382, 1317, 1200. δH (500 MHz, CF3COOD)
10H), 3.94 (br t, 2H), 4.93 (spt, 1H, J = 6.0), 5.77 (m, 2H), 7.74 2.58 (s, 6H), 2.60 (br s, 4H), 2.90 (br s, 2H), 3.19 (br s, 4H), 3.61
(s, 2H), 7.88 (s, 1H), 8.13 (br t, 1H), 8.25 (br d, 1H), 8.48 (br t, (t, 4H, J = 7.0), 3.80 (br t, 2H), 4.46 (br t, 4H), 5.82 (br t, 2H),
1H), 8.69 (br d, 2H), 8.74 (d, 1H, J = 9.0). MALDI-TOF MS m/z 7.83 (br s, 2H), 8.08 (s, 2H), 8.14 (d, 2H, J = 9.0), 8.81 (d, 2H,
1204.8 M+; HRMS m/z 1204.4670 M+ (1204.4616 calculated).
J = 9.0). MALDI-TOF MS m/z 866.8 (M + H)+, HRMS m/z
Compound 12. A mixture of compound 1014 (30 mg, 867.1417 M+ (867.1411 calculated).
0.072 mmole), distilled 1,3-propane sultone (0.132 g,
1.08 mmole, 15 equivalents) and anhydrous sodium carbonate
(38 mg, 0.36 mmole, 5 equivalents) in [BMIM][PF6] (1 mL) was
2. Synthesis of anti-HBsAg antibody conjugates of 9, 13 and
14 (Table 1)
heated at 150 °C under a nitrogen atmosphere. After 3 hours, A murine anti-HBsAg monoclonal antibody with a neutral pI
an additional 15 equivalents of 1,3-propane sultone and 5 was used for labeling with the acridinium ester labels 9, 13
equivalents of sodium carbonate were added and the reaction and 14.
was continued at 150 °C for 3 hours. A final addition of 15
The monoclonal antibody (1 mg, 6.67 nanomoles, stock
equivalents of 1,3-propane sultone and 5 equivalents of solution 6.2 mg mL−1, 0.161 mL) was diluted with 0.24 mL of
sodium carbonate was then performed and the reaction was 0.1 M sodium carbonate, pH = 9. The protein solution was
continued at 150 °C for 3 hours (total reaction time 9 hours). treated with DMSO solutions of acridinium esters as follows:
The reaction was then cooled to room temperature and ana- for labeling with 5 and 10 equivalents of 14, 0.0068 mL and
lyzed by HPLC using a Phenomenex, C18, 10 micron, 3.9 × 0.0136 mL of a 5 mg mL−1 DMSO solution were added; for
300 mm column and a 30 minute gradient of 10 → 70% labeling with 5 and 10 equivalents of 9, 0.0081 and 0.0162 mL
MeCN–water (each with 0.05% TFA) at a flow rate of 1.0 mL of a 5 mg mL−1 DMSO solution were added; and, for labeling
per minute and UV detection at 260 nm. Compound 11 was with 5 and 10 equivalents of 13, 0.0058 mL and 0.0116 mL of a
observed eluting as a broad peak at 14 minutes. The crude 5 mg mL−1 DMSO solution were added. The labeling reactions
reaction mixture was partitioned between ethyl acetate (20 mL) were stirred at 4 °C for ∼3 hours and were then transferred to
and water (20 mL). The dark brown aqueous layer containing 4 mL Amicon™ filters (MW 30 000 cutoff) and diluted with
product was separated and washed once with ethyl acetate 3.6 mL de-ionized water. The volume was reduced to ∼0.25 mL
(20 mL). It was then concentrated under reduced pressure. by centrifuging at 4000g for 10 minutes. The concentrated con-
Crude 11 was suspended in 1 M HCl (20 mL) and refluxed for jugate solutions were diluted with 4 mL de-ionized water and
4 hours. It was then cooled to room temperature and analyzed centrifuged again to reduce the volume. This process was
by HPLC which showed complete conversion to product 12 repeated a total of four times. Finally, the concentrated conju-
eluting at 12.4 minutes. HPLC analysis of the reaction mixture gates were diluted with ∼0.25 mL de-ionized water and were
using a Phenomenex C8, 5 micron, 4.6 × 100 mm column and frozen and lyophilized. The lyophilized conjugates were dis-
a 40 minute gradient of 10 → 60% MeCN–water (each with solved in 0.5 mL of de-ionized water. Label incorporation was
0.05% TFA) at a flow rate of 1 mL per minute showed product measured by MALDI-TOF mass spectrometry using the
12 eluting at 8.5 minutes. The product was purified by prepara- Voyager-DE instrument from Perkin-Elmer. This entailed
tive HPLC as described for compound 2. The HPLC fractions measuring the molecular weight of the unlabeled protein and
containing product were frozen at −80 °C and lyophilized. the labeled protein. The acridinium ester label contributed to
Yield = 30 mg (55%); dark yellow powder. νmax/cm−1 3477 the observed difference in mass between these two measure-
(OH), 1751 and 1719 (CO), 1615, 1465, 1381, 1224, 1145. δH ments. By knowing the molecular weight of the specific acridi-
(500 MHz, CF3COOD) 2.51 (s, 6H), 2.54 (br t, 4H), 2.84 (br s, nium ester label, the extent of label incorporation for that
2H), 3.57 (t, 4H, J = 7.0), 3.74 (br t, 2H), 4.40 (t, 4H, J = 5.0), specific acridinium ester could thus be calculated. Label incor-
5.76 (m, 2H), 7.78 (br s, 2H), 8.01 (s, 2H), 8.07 (d, 2H, J = 10.0), poration in each conjugate is tabulated in Table 1.
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1037

View Article Online
Paper
Organic & Biomolecular Chemistry
3. Emission wavelength measurements (Fig. 4)
chemiluminescence reaction was carried out a minimum of
three times, averaged and converted to a percentage of the che-
miluminescence accumulated up to each time interval. The
output from the luminometer instrument was expressed as
RLUs (relative light units).
Emission spectra of the acridinium esters 9, 13 and 14 were
measured using an FSSS (Fast Spectral Scanning System)
camera (Spectra Scan Model 704) from Photo Research Inc. In
a typical measurement, 0.05–0.1 mL of a 1 mg mL−1 solution
of the acridinium ester in a 2 : 1 mixture of water–MeCN (with
0.05% TFA) was diluted with 0.3 mL of reagent 1 comprising
0.5% hydrogen peroxide in 0.1 M nitric acid. Just prior to the
addition of reagent 2 comprising 7 mM CTAC in 0.25 M
sodium hydroxide, the shutter of the camera was opened and
light was collected for 5 seconds. The output of the instrument
is a graph of light intensity versus wavelength.
6. Isoelectric focusing (IEF) gel electrophoresis
Isoelectric focusing (IEF) gel electrophoresis of the anti-HBsAg
antibody and the acridinium ester conjugates was performed
using an XCell SureLock Mini-Cell (Life Technologies) and
Novex 3-10 IEF gels (Life Technologies) and Novex buffers. IEF
markers, pH 3–10 (Life Technologies), were used as reference
standards. Conjugates (∼2 mg mL−1) were mixed 1 : 1 with
Novex IEF sample buffers pH 3–10 and 0.012 mL was loaded
onto the gel. The gel was run at 100 volts for 1 hour, 200 volts
for 1 hour and then 500 volts for 30 minutes. The gel was then
fixed for 30 minutes using 12% trichloroacetic acid and 3.5%
sulfosalicylic acid. The gel was stained for 15 minutes in
Coomassie Blue R-250 and then doubly stained overnight
with 30% methanol, 10% acetic acid in de-ionized water.
4. UV-Visible spectrophotometric measurements (Fig. 5)
UV-Visible spectrophotometry of acridinium esters 9, 13 and
14 for determination of the pKa of the acridinium to pseudo-
base transition was carried out by dissolving the HPLC-puri-
fied acridinium esters in DMSO (∼1 mg mL−1). These
solutions were further diluted 20-fold into 0.2 M phosphate or
borate buffer at pH 2, 3.0, 4.0, 5.0, 6.0, 7.0, 8.0, 9.0 and 10.0
(8–10 only for compound 13, pH 10 was 0.2 M borate buffer). A 7. Measurement of fractional non-specific binding (FNSB)
solution of 0.1 M HCl was used for pH 1. The diluted acridi- (Table 2)
nium ester solutions were allowed to stand for one hour at
The fractional non-specific binding (FNSB) of anti-HBsAg anti-
room temperature (9 and 14) or 16 hours (13) and then UV-Vi-
body conjugates of acridinium ester labels 9, 13 and 14 was
sible spectra were recorded from 220–500 nm. The absorption
measured using a Siemens Healthcare Diagnostics’ ADVIA
intensity of the acridinium band at 383 nm for compounds 9,
Centaur® automated immunoanalyzer. The conjugates were
diluted to equivalent concentrations of 10 nM in a solution
consisting of 0.1 M sodium N-(2-hydroxyethyl) piperazine-N′-2-
14 and 410 nm for compound 13 was measured as a function
of pH. A plot of this data is shown in Fig. 5.
ethanesulfonate (HEPES), 0.15 M sodium chloride, 7.7 mM
sodium azide, 12 mM t-octylphenoxypolyethoxyethanol (Triton
5. Chemiluminescence measurements (Fig. 6, 7, S10†)
Chemiluminescence of acridinium esters 9, 13 and 14 and X-100), 0.076 mM BSA, pH = 7.7. A volume of 0.1 mL each of
their anti-HBsAg antibody conjugates was measured on an these 10 nM protein–acridinium ester conjugate solutions
AutoLumat LB953 Plus luminometer from Berthold Techno- were mixed with 0.1 mL of horse serum and 0.2 mL of either
logies. HPLC-purified acridinium esters 9, 13 and 14 were of three solid phases. The first solid phase was 0.35 grams per
initially dissolved in DMSO (1–2 mg mL−1), allowed to stand liter paramagnetic microparticles (PMP) covalently covered
for several hours and were further diluted for chemilumines- with mouse anti-fluorescein antibody. The second solid phase
cence measurements in an aqueous buffer of 10 mM disodium was 0.35 grams per liter of Dynabeads® M-280 magnetic latex
hydrogen phosphate, 0.15 M NaCl, 8 mM sodium azide and particles (Life Technologies) covalently covered with streptavi-
0.015 mM bovine serum albumin (BSA), pH = 8.0. Protein con- din. The third solid phase was 0.35 grams per liter of Dyna-
jugates, ∼2 mg mL−1 were serially diluted 105-fold for chemilu- beads® M-270 magnetic latex particles (Life Technologies)
minescence measurements. Similarly, solutions of acridinium covalently covered with streptavidin. After an incubation of
ester labels were serially diluted 106-fold for chemilumines- 5.5 minutes to allow interaction between the acridinium ester
cence measurements. A 0.010 mL volume of each diluted acri- conjugate and the solid phases, the solid phases were magneti-
dinium ester or conjugate sample was dispensed into the cally collected and washed twice with either water or a buffer
bottom of a cuvette. Cuvettes were placed in the primed LB953 of 10 mM potassium phosphate, 0.14 M sodium chloride,
and the chemiluminescence reaction was initiated with the 14 mM sodium azide, 1 mM EDTA (ethylenediaminetetraacetic
sequential addition of 0.3 mL of reagent 1, a solution of 0.5% acid), 0.8 mM Tween 20 (polyoxyethylene-20 sorbitan mono-
hydrogen peroxide in 0.1 M nitric acid, followed by the laurate), pH = 7.2. The chemiluminescence, expressed as
addition of 0.3 mL of reagent 2, a solution of 0.25 M sodium RLUs, of both the input acridinium ester chemiluminescence
hydroxide with or without CTAC surfactant. Each chemilumi- and the chemiluminescence of the acridinium ester associated
nescence flash curve was measured in 100 intervals of 0.1 with the particles was measured for 4 seconds on the ADVIA
second (10 seconds total time) in the presence of 7 mM CTAC Centaur® with sequential addition of 0.3 mL each of reagent 1
or 240 intervals of 0.5 second (2 minutes total time) in the (0.1 M nitric acid and 0.5% hydrogen peroxide) and reagent 2
absence of CTAC from the point of chemiluminescence (0.25 M sodium hydroxide containing 7 mM cetyltrimethylam-
initiation with the addition of 0.25
M
NaOH. Each monium chloride). FNSB was calculated as the ratio of
1038 | Org. Biomol. Chem., 2013, 11, 1026–1039
This journal is © The Royal Society of Chemistry 2013

View Article Online
Organic & Biomolecular Chemistry
Paper
particle-bound chemiluminescence to input chemilumines-
cence by averaging the chemiluminescence of ten
measurements.
Y. Goda, Can. J. Chem., 1984, 62, 351–354;
(c) P. W. Hammond, W. A. Wiese, A. A. Waldrop III,
N. C. Nelson and L. J. Arnold Jr., J. Biolumin. Chemilumin.,
1991, 6, 35–43.
6 A. Natrajan, T. Sells, H. Schroeder, G. Yang, D. Sharpe,
Q. Jiang, H. Lukinsky and S.-J. Law, US Pat, 7,319,041,
2008.
7 (a) T. Kinkel, H. Lűbbers, E. Schmidt, P. Molz and
H. J. Skrzipczyk, J. Biolumin. Chemilumin., 1989, 4, 136–139;
(b) P. G. Mattingly, J. Biolumin. Chemilumin., 1991, 6, 107–114.
8 S.-J. Law, T. Miller, U. Piran, C. Klukas, S. Chang and
J. Unger, J. Biolumin. Chemilumin., 1989, 4, 88–98.
9 A. Natrajan and D. Wen, Green Chem., 2011, 13, 913–921.
8. Measurement of chemiluminescence stability (Fig. 9)
Acridinium ester anti-HBsAg antibody conjugates of labels 9,
13 and 14 were diluted to a concentration of 0.1 nM in a buffer
of 0.1 M sodium N-(2-hydroxyethyl) piperazine-N′-2-ethane-
sulfonate (HEPES), 0.15 M sodium chloride, 7.7 mM sodium
azide, 12 mM t-octylphenoxypolyethoxyethanol (Triton X-100),
0.076 mM BSA at pH = 6.0 or pH 7.4. The diluted conjugates
were kept at 4 and 37 °C for four weeks. The residual chemi-
luminescence of each diluted conjugate was assessed over the
course of the four weeks by periodically averaging the 5 second 10 R. C. Brown, Z. Li, A. J. Rutter, X. Mu, O. H. Weeks,
chemiluminescence measurement of five 0.01 mL samples on
the AutoLumat LB953 Plus luminometer with sequential
K. Smith and I. Weeks, Org. Biomol. Chem., 2009, 7, 386–
394.
addition of 0.3 mL each of reagent 1 (0.1 M nitric acid and 11 S. Caddick, J. D. Wilden and D. B. Judd, J. Am. Chem. Soc.,
0.5% hydrogen peroxide) and reagent 2 (0.25 M sodium 2004, 126, 1024–1025.
hydroxide containing 7 mM cetyltrimethylammonium chlor- 12 J. D. Wilden, D. B. Judd and S. Caddick, Tetrahedron Lett.,
ide). The change in chemiluminescence was gauged against
initial values as a percentage.
2005, 46, 7637–7640.
13 (a) A. T. Ślósarczyk and L. Baltzer, J. Pept. Sci., 2012, 18,
261–269; (b) A. T. Ślósarczyk, R. Ramapanicker, T. Norberg
and L. Baltzer, RSC Adv., 2012, 2, 908–914.
Acknowledgements
14 A. Natrajan, Q. Jiang, D. Sharpe and J. Costello, US Pat.,
7,309,615 B2, 2007.
We thank Ms Elzbieta Rotter-Wask for the IEF gel of the anti-
body conjugates. We also thank Mr David Wen for IR spectra
of synthetic compounds and for helpful suggestions with the
manuscript.
15 K. A. Browne, D. D. Deheyn, G. A. El-Hiti, K. Smith and
I. Weeks, J. Am. Chem. Soc., 2011, 133, 14637–14648.
16 (a) F. McCapra, J. Chem. Soc., Chem. Commun., 1977, 946–
948; (b) F. McCapra, I. Beheshti, A. Burford, R. A. Hann
and K. A. Zaklika, J. Chem. Soc., Chem. Commun., 1977,
944–946.
References
17 G. B. Schuster, Acc. Chem. Res., 1979, 12, 366–373.
1 (a) S.-J. Law, C. S. Leventis, A. Natrajan, Q. Jiang, 18 D. E. Garfin, in Gel Electrophoresis of Proteins, Essential Cell
P. B. Connolly, J. P. Kilroy, C. R. McCudden and
S. M. Tirrell, US Pat., 5,656,426, 1997; (b) A. Natrajan,
D. Sharpe and Q. Jiang, US Pat., 6,664,043 B2, 2003;
Biology, Volume 1, Cell Structure: A Practical Approach, ed.
J. Davey and M. Lord, Oxford University Press, 2003.
pp. 197–268.
(c) A. Natrajan, D. Sharpe, J. Costello and Q. Jiang, Anal. 19 (a) R. C. Chapman, E. Ostuni, L. Yan and G. M. Whitesides,
Biochem., 2010, 406, 204–213; (d) A. Natrajan, D. Sharpe
and D. Wen, Org. Biomol. Chem., 2012, 10, 1883–1895.
2 (a) F. McCapra, Acc. Chem. Res., 1976, 9/6, 201–208;
(b) F. McCapra, D. Watmore, F. Sumun, A. Patel,
I. Beheshti, K. Ramakrishnan and J. Branson, J. Biolumin.
Chemilumin., 1989, 4, 51–58; (c) J. Rak, P. Skurski and
J. Błażejowski, J. Org. Chem., 1999, 64, 3002–3008;
(d) K. Krzymiński, A. Ożóg, P. Malecha, A. D. Roshal,
A. Wróblewska, B. Zadykowicz and J. Błażejowski, J. Org.
Chem., 2011, 76, 1072–1085.
3 A. Natrajan, D. Sharpe and D. Wen, Org. Biomol. Chem.,
2011, 9, 5092–5103.
4 A. Natrajan, D. Sharpe and D. Wen, Org. Biomol. Chem.,
2012, 10, 3432–3447.
5 (a) I. Weeks, I. Behesti, F. McCapra, A. K. Campbell and
J. S. Woodhead, Clin. Chem., 1983, 29/8, 1474–1479;
(b) J. W. Bunting, V. S. F. Chew, S. B. Abhyankar and
Langmuir, 2000, 16, 6927–6936; (b) R. C. Chapman,
E. Ostuni, M. N. Liang, G. Meluleni, E. Kim, L. Yan, G. Pier,
H. S. Warren and G. M. Whitesides, Langmuir, 2001, 17,
1225–1233; (c) S. J. Sofia, V. Premnath and E. W. Merrill,
Macromolecules, 1998, 31, 5059–5070; (d) Y.-H. Zhao,
B.-K. Zhu, L. Kong and Y.-Y. Xu, Langmuir, 2007, 23, 5779–
5786; (e) A. R. Denes, E. B. Somers, A. C. L. Wong and
F. Denes, J. Appl. Polym. Sci., 2001, 81, 3425–3438;
(f) J. Ladd, Z. Zhang, S. Chen, J. C. Hower and S. Jiang, Bio-
macromolecules, 2008, 9, 1357–1361; (g) Y. Chang, S. Chen,
Z. Zhang and S. Jiang, Langmuir, 2006, 22, 2222–2226;
(h) Z. Zhang, T. Chao, S. Chen and S. Jiang, Langmuir, 2006,
22, 10072–10077; (i) W. K. Cho, B. Kong and I. S. Choi,
Langmuir, 2007, 23, 5678–5682; ( j) G. Cheng, Z. Zhang,
S. Chen, J. D. Bryers and S. Jiang, Biomaterials, 2007, 28,
4192–4199; (k) H. Kitano, A. Kawasaki, H. Kawasaki and
S. Morokoshi, J. Colloid Interface Sci., 2005, 282, 340–348.
This journal is © The Royal Society of Chemistry 2013
Org. Biomol. Chem., 2013, 11, 1026–1039 | 1039