Photoinduced DNA cleavage and cellular damage in human dermal fibroblasts by 2,3-diaminophenazine

Article abstract of DOI:10.1562/2004-07-20-RA-237.1

Aromatic amines, such as o-phenylenediamine (OPD), have been used extensively in commercial hair dyes and in the synthesis of agricultural pesticides. Air oxidation of OPD resells in the formation of 2,3-diaminophenazine (DAP). Although the mutagenic toxicity of DAP has been shown in both prokaryotic and eukaryotic systems, its phototoxicity remains largely unexplored. This study focuses on the pH-dependent photophysical properties of DAP and demonstrates its ability to photoinduce DNA damage to pUC19 plasmid in vitro. The photocytotoxicity of DAP toward human skin fibroblasts was also measured. DAP exhibits weak intercalative binding to double-stranded DNA with a binding constant K_b = 3.5 × 10 ³ M^-1. Furthermore, upon irradiation with visible light, DAP is able to nick plasmid DNA in the presence of oxygen. The concentration of DAP that resulted in 50% cell death was 172 ± 9 μM in the dark and 13 ± 1 μM after irradiation of the DAP-treated cell cultures with visible light (400-700 nm, 30 min, 5 J/ cm²). The 13-fold increase in toxicity upon exposure to visible light shows the need for further study of the photocytotoxicity of contaminants such as DAP.

Full text of DOI:10.1562/2004-07-20-RA-237.1

Photoinduced DNA Cleavage and Cellular Damage in Human Dermal Fibroblasts
by 2,3-Diaminophenazine
Author(s): Patty K-L. Fu, Sonia Abuzakhm, Claudia Turro
Source: Photochemistry and Photobiology, 81(1):89-95.
Published By: American Society for Photobiology
DOI: http://dx.doi.org/10.1562/2004-07-20-RA-237.1
URL: http://www.bioone.org/doi/full/10.1562/2004-07-20-RA-237.1
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Photochemistry and Photobiology, 2005, 81: 89–95
Photoinduced DNA Cleavage and Cellular Damage in Human Dermal
Fibroblasts by 2,3-Diaminophenazine{
Patty K.-L. Fu*1, Sonia Abuzakhm2 and Claudia Turro*2
1Center for Food Safety and Applied Nutrition, U.S. Food and Drug Administration, College Park, MD
2Department of Chemistry, The Ohio State University, Columbus, OH
Received 20 July 2004; accepted 7 October 2004
in the ELISA assays permits the use of spectrofluorometric
detection, which results in a significant increase in sensitivity (5–7).
ABSTRACT
Aromatic amines, such as o-phenylenediamine (OPD), have
been used extensively in commercial hair dyes and in the
synthesis of agricultural pesticides. Air oxidation of OPD
results in the formation of 2,3-diaminophenazine (DAP).
Although the mutagenic toxicity of DAP has been shown in
both prokaryotic and eukaryotic systems, its phototoxicity
remains largely unexplored. This study focuses on the pH-
dependent photophysical properties of DAP and demonstrates
its ability to photoinduce DNA damage to pUC19 plasmid
in vitro. The photocytotoxicity of DAP toward human skin
fibroblasts was also measured. DAP exhibits weak intercalative
binding to double-stranded DNA with a binding constant Kb 5
3.53103 M21. Furthermore, upon irradiation with visible light,
DAP is able to nick plasmid DNA in the presence of oxygen. The
concentration of DAP that resulted in 50% cell death was 172 6
9 lM in the dark and 13 6 1 lM after irradiation of the DAP-
treated cell cultures with visible light (400–700 nm, 30 min, 5 J/
cm2). The 13-fold increase in toxicity upon exposure to visible
light shows the need for further study of the photocytotoxicity
of contaminants such as DAP.
Two toxic byproducts formed upon routine handling of OPD are
2,3-diaminophenazine (DAP) and 2-amino-3-hydroxyphenazine
(AHP) (1–13). DAP is considered to be the major mutagenic
contaminant generated from the oxidation of OPD (Fig. 1), which
can be carried out by oxygen in air and is accelerated by exposure to
light (1–15). AHP results from a chemical transformation of DAP in
aqueous media (12,13). Both DAP and AHP are well-recognized
toxic environmental contaminants and industrial occupational
hazards (1,2,4,12,13). The mutagenic effects and the genotoxicity
of DAP and AHP have been reported in both prokaryotic and
eukaryotic systems (12–14), and it has been shown that DAP
induces chromosomal DNA damage in bacteria, human blood
lymphocytes and mice (1,2,12–15).
The photophysical properties of DAP remain largely unexplored,
although the molecule is known to be highly emissive (5–7,16–22).
Most investigations of DAP fluorescence have typically focused on
its use in the development of new analytical applications and
detection techniques (5–7,16–22); however, the potential biological
impact of its excited-state photochemistry remains unknown.
Because genotoxicity tests to date have focused on the ground-
state reactivity (1–4,12–15) and were, therefore, carried out in the
absence of irradiation, the photobiological effects and the photo-
toxicity of DAP are not well understood. Because of the potential
human exposure to DAP generated from the oxidation of OPD in
topically applied hair dyes, it is important to further examine the
excited-state properties and reactivity of DAP toward biomolecules.
In this study, we explore the photophysical and photochemical
properties of DAP, as well as its ability to sensitize light-induced
DNA cleavage in plasmid, and cellular damage in human skin
fibroblasts.
INTRODUCTION
Phenylenediamines, such as o-phenylenediamine (OPD), are the
most common and widely used aromatic amines for the formulation
of commercial hair dyes and synthesis of agricultural fungicides (1–
4). In addition, OPD is one of the most effective chromogenic
substrates for a variety of enzyme-linked immunosorbent assays
(ELISA) used in the study of enzyme kinetics (5–7). The use of OPD
{Posted on the website on 19 October 2004.
MATERIALS AND METHODS
*To whom correspondence should be addressed (PKF) at: Center for Food
Safety and Applied Nutrition, U.S. Food and Drug Administration, 5100
Paint Branch Parkway, College Park, MD 20740, USA. Fax: 301-436-
2897; e-mail: patty.fu@FDA.GOV
Materials and chemicals. FeCl3, OPD, HCl, D2O, ethylenediaminetetra-
acetic acid (EDTA) and formamide were purchased from Aldrich (St.
Louis, MO). Ethidium bromide (EtBr), NaCl, calf thymus DNA (CT DNA),
dialysis membrane (.100 000 Da), Tris–acetate EDTA (TAE) buffer,
Na2HPO4, NaH2PO4, agrose, N-lauroyl sarcosine, micro-Lowry reagent kit
690-A and Folin–Ciocalteau phenol reagent were purchased from Sigma
(St. Louis, MO). Sonicated herring sperm DNA purchased from Promega
(Madison, WI). pUC19 plasmid and gel loading solution were purchased
from New England Biolabs (Beverly, MA). Dulbecco modified Eagle
media, fetal bovine serum, phosphate-buffered saline (PBS), gentamicin,
glucose, L-glutamine were purchased from Invitrogen Corp (Carlsbad, CA).
Synthesis. DAP was synthesized from FeCl3 and OPD according to
a reported method (23). FeCl3 (0.7120 g) was dissolved in 20 mL of H2O
*To whom correspondence should be addressed (CT) at: Department of
Chemistry, The Ohio State University, Columbus, OH 43210, USA. Fax:
614-292-1685; e-mail: turro@chemistry.ohio-state.edu
Abbreviations: AHP, 2-amino-3-hydroxyphenazine; CT DNA, calf thymus
DNA; DAP, 2,3-diaminophenazine; ds-DNA, double-stranded DNA;
EDTA, ethylenediaminetetraacetic acid; ELISA, enzyme-linked immu-
nosorbent assay; EtBr, ethidium bromide; 1H NMR, proton nuclear
magnetic resonance; LC50, lethal concentration 50; OPD, o-phenylenedi-
amine; PBS, phosphate-buffered saline; TAE, Tris–acetate.
Ó 2005 American Society for Photobiology 0031-8655/05
89

90 Patty K.-L. Fu et al.
Spectrophotometric titration. The binding constant of DAP to CT DNA
was determined by absorption titration at room temperature using 10
individual freshly prepared solutions with a constant concentration of DAP
(25 lM) and CT DNA concentrations that ranged from 0 to 125 lM (5 mM
NaCl, 10 mM phosphate buffer, pH 7.5). The binding constant, Kb, was
determined from the absorption changes during the DNA titration and was
calculated using the equation (ea ꢀ eb)/(eb ꢀ ef) 5 (1/Kb) 3 (1/[DNA]) þ 1,
where ea, ef and eb are the molar extinction coefficients for the apparent
absorption of the complex at a given DNA concentration, for the complex
free in solution and for the complex fully bound to DNA, respectively. The
value of eb was determined from the titration where further addition of
DNA did not result in changes to the spectrum. The binding constant for
each molecule was determined by plotting (ea ꢀ eb)/(eb ꢀ ef) vs 1/[DNA],
and Kb was obtained as the reciprocal value of the slope (28,29).
Viscosity measurements. The viscosity experiments were performed using
a Cannon-Manning semi-micro size 75 viscometer (State College, PA)
immersed in a thermostated water bath maintained at 258C (60.1). Solutions
(1 mL 25 mM phosphate buffer, pH 5 7.2) containing 2.5 mM (base pairs)
sonicated herring sperm DNA purchased from Promega and used without
further purification. Titrations were conducted by the successive addition of
either EtBr or DAP directly into the viscometer without removing the DNA
solution, and the solutions were bubbled with nitrogen after each addition to
ensure mixing (30–32). The data points were plotted as (g/g0)1/3 vs
[Compound]/[DNA bp], where g0 and g represent the viscosity of the DNA
solution in the absence and presence of a given compound, respectively, both
relative to the viscosity of the buffer at the same temperature (30–33). Both g
and g0 were calculated from the observed flow times, t, relative to the flow
time for the buffer, t0 (98.5 6 0.5 s). The flow time of DNA alone was
measured to be 140.2 6 0.5 s at 258C.
DNA photocleavage. The DNA photocleavage experiments were carried
out on a 20 lL total sample volume in 0.5 mL transparent eppendorf
microtubes containing 120 lM pUC19 plasmid and 30 lM of DAP in 10
mM phosphate buffer, 5 mM NaCl, pH 5 7.5. All samples were irradiated
for 30 min after 30 min of incubation following addition of DAP. After
irradiation, the samples were loaded into the gel following the addition of
4 lL of DNA gel loading solution to each reaction mixture. In determining
the pH dependence of photocleavage, the pH of each reaction mixture was
varied using phosphate buffer before adding DAP and irradiation. All the
photolysis experiments were conducted in air at room temperature. The
electrophoresis was carried out using 2% agarose gel stained with 0.5 mg/L
EtBr in 13 TAE running buffer at 90 V for 1.5 h and the gels were imaged
on a GelDoc 2000 transilluminator (Bio-Rad, Hercules, CA). The extent of
photocleavage was calculated from the ratio of nicked (circular) to
supercoiled double-stranded DNA (ds-DNA) determined from the in-
tegration of image intensity available in the QuantityOne Analysis System
software (Bio-Rad). In these measurements, no correction was made for the
difference in EtBr emission from the different forms of DNA.
Cytotoxicity and photocytotoxicity assays. Human skin fibroblasts
(Hs-27) were obtained from the American Type Culture Collection, cell
line CRL-1634 (Manassas, VA). The cells were cultured in Dulbecco
modified Eagle media, containing 10% fetal bovine serum, 50 lg/mL
gentamicin, 4.5 mg/mL glucose and 4 mM L-glutamine. The cell cultures
were incubated in a humidified atmosphere containing 5% CO2 at 378C.
For assessing the cytotoxicity and photocytotoxicity of DAP, .80%
confluent monolayers of Hs-27 in 60 mm culture dishes were used. The
monolayers were first washed twice with PBS to ensure that the culture
dishes were free of any culture medium. Fresh PBS (3 mL) containing
various concentrations of DAP was then added to cover the fibroblasts. The
cells were irradiated through the PBS buffer, which does not absorb light in
the visible region, immediately without an incubation period. The spectral
output of the visible light was ;2.8 3 10ꢀ3 W/cm2, and all samples were
irradiated no longer than 30 min (;5 J/cm2) at room temperature. After
irradiation, the cells were removed from the dishes by trypsinization,
reseeded into 24-well culture dishes and incubated for 48 h. N-lauroyl
sarcosine (200 lL, 40 mM) was then added to each well and the cells were
allowed to lyse for at least 30 min. Quantitative determination of the protein
content in each well was undertaken using Peterson modification of the
micro-Lowry method. The lysate was treated with 200 lL Lowry reagent
for 30 min and then with 100 lL Folin–Ciocalteau phenol reagent for 30
min. A portion (200 lL) of the contents of each well was transferred to
a 96-well plate for absorbance determination using a 96-well MRX plate
reader (Dynatech Laboratory, DynPort, VA). The absorbance measured at
630 nm is proportional to the total protein content and the number of cells
in each well.
Figure 1. Structure of (a) OPD, (b) phenazine, (c) DAP, (d) acridine ring
and (e) proflavin.
and was poured into a stirring solution containing OPD (0.2160 g) and 20
mL of HCl in 0.6 mL H2O at room temperature. The red precipitate was
filtered and washed repeatedly with diethylether and methanol. Proton
nuclear magnetic resonance (1H NMR) (400 MHz, D2O): d 5 8.0 (m, 2H);
7.7 (m, 2H); 6.9 (s, 2H); 6.2 (s, 4H). The 1H NMR spectrum of the product
in D2O was consistent with that reported previously for DAP (23).
Absorption, fluorescence and irradiation source. Absorption measure-
ments were performed using a Hewlett Packard diode array spectrometer
(HP8453) with HP8453Win System software installed on an HP Vectra XM
5/120 desktop computer (Loveland, CO). Emission spectra for the pH
titrations were collected using a SPEX FluoroMax-2 spectrometer (Edison,
NJ) equipped with
a 150 W Xenon source, a red-sensitive 928P
photomultiplier tube and DataMax-Std software on a Pentium microproces-
sor. The emission experiments in the presence of DNA (pH57.5 with kexc5
430 nm and kem5590 nm) were carried out using a MFX fluorimeter (Dynex
Technology, Chantilly, VA) using Revelation software on a Micron PC. The
source of visible radiation (400–700 nm) for DNA photocleavage and
photocytotoxicity was a photoreactor (Luzchem, Ontario, Canada) com-
posed of twelve 8 W GE watt-miser bulbs (Luzchem). The light intensity was
measured using a radiometer (International Light Inc., Newburyport, MA).
Equilibrium dialysis. CT DNA was purchased from Sigma and purified
using a standard literature procedure (24). First, the same volume of
formamide denaturing buffer (20 mM Tris–HCl, pH 8.0, 0.8 M NaCl, 80%
vol/vol formamide) was added to the CT DNA solution (1 mg/mL) and left
for 12 h at 158C, which results in the denaturation of any proteins bound to
CT DNA. After the incubation, the solution was dialyzed using .100 000 Da
dialysis membrane (Sigma) for 45 min against 1 L of a buffer containing 20
mM phosphate buffer (pH58.0), 0.1 M NaCl and 10 mM EDTA; the buffer
was replaced and the dialysis repeated three times in 4 h intervals. The CT
DNA was then dialyzed twice against 10 mM phosphate buffer (pH 8.0), 10
mM NaCl, 0.5 mM EDTA for 8–10 h. The dialysis membrane was prepared
according to standard methods (24). The 10–20 cm long pieces of tubing
were boiled for 10 min in ;500 mL of 2% (wt/vol) sodium bicarbonate and 1
mM EDTA (pH58.0),were then rinsed thoroughly with deionized H2O and
were then boiled for 10 min in 1 mM EDTA and allowed to cool. The tubes
were stored at 48C submerged in 1 mM EDTA and were rinsed thoroughly
with deionized H2O before use.
In the binding constant determination using dialysis, the total con-
centration of DAP (Ct) present in the dialysate solution was determined
spectrophotometrically using an extinction coefficient, e, of 13 000 cmꢀ1
M
ꢀ1 at 440 nm. The exctinction coefficent was measured using Beer’s Law.
CT DNA (100 lM) was placed in a dialysis membrane and was dialyzed
against 1.0 L of 10 lM DAP in circulating buffer (6 mM Na2HPO4, 3 mM
NaH2PO4, 1 mM EDTA and 125 mM NaCl, pH 5 7.0) for 48 h. The free
DAP concentration (Cf) was determined from an aliquot of the dialysate
solution after the 48 h equilibration period. The amount of DAP bound to
the CT DNA, Cb, was determined from the mass balance equation given by
Cb 5 Ct ꢀ Cf (25–27).

Photochemistry and Photobiology, 2005, 81 91
RESULTS AND DISCUSSION
Photophysical properties in solution
The absorption spectrum of DAP in H2O is characterized by
a maximum at 420 nm (e513 000 Mꢀ1 cmꢀ1), and excitation in the
visible region (400–450 nm) results in emission with a maximum at
583 nm. Pronounced changes in the electronic absorption and
emission spectra of DAP as a function of pH are observed and are
shown in Fig. 2. The absorption maximum of DAP shifts from 450
to 417 nm as the pH is varied from 4.0 to 8.0 (Fig. 2a). Weak
emission is observed at pH 5 3.0 with a maximum at 575 nm, and
a 13-fold increase in intensity is observed as the pH is raised to 9.0
with a shift in the maximum to 585 nm (Fig. 2b). The pH-dependent
changes we observe in the absorption and the emission intensities
result from two protonation steps at pH 5 5 and pH 56.6, agreeing
with the previously published pka value of 5.1 (34). The changes in
absorption and emission as a function of solution pH are in-
dicative of the existence of ground-state species with different
degrees of protonation, which possess different luminescence pro-
perties (35–44).
The pH and solvent dependence of the emission of various
heterocyclic aromatic dyes, such as acridine, phenazine, proflavin
(3,6-diaminoacridine) and their derivatives (Fig. 1) have been
investigated previously (35,36,40). The redshift of the absorption
maximum of DAP shown in Fig. 2a as the solution pH is decreased
is similar to that observed in proflavin. This pH-dependent redshift
was attributed to the protonation of proflavin’s ring nitrogen with
a pka 5 9.5 (35,36). The protonation of the amino nitrogen in
proflavin with pka values of 0.2 does not result in significant
spectral changes (35,40). The protonation of the nitrogen in the
aromatic ring of phenazine and acridine with pka values of 1.21 and
5.4, respectively, has also been reported to result in significant
spectral changes both in the absorption and emission maxima
(39,42,43). Because amination increases the basicity of the
aromatic nitrogen in acridine, the amino substituents in DAP
result in a shift in the pka value from 1.21 in phenazine to 5.1 for
DAP. The magnitude of this shift is similar to that observed for
protonation of the aromatic nitrogen between acridine (pka 5 5.4)
and proflavin (pka 5 9.5). Therefore, it is likely that the observed
spectral shifts and intensity changes in DAP are because of
protonation of the nitrogens in the aromatic ring.
Figure 2. Room temperature (a) electronic absorption and (b) emission
(kexc 5 440 nm) spectra of DAP at different pH values.
polymerization because of exposure to low levels of ultraviolet–
visible light (45–47), 10 individual solutions were freshly prepared
with a constant concentration of DAP (25 lM) and ds-DNA
concentrations that ranged from 0 to 125 lM. The absorption
spectrum of 25 lM DAP showed a notable 20 6 1% hypochromicity
as ds-DNA concentration was increased to 125 lM, along with
a small redshift of the absorption maximum (;3 nm). A binding
constant Kb52.53103 Mꢀ1 was obtained using the changes in DAP
absorption at 420 nm as a function of ds-DNA concentration at
pH 5 7.0 by methods reported previously (28,29,48–50). A slight
decrease in Kb (2.1 3 103
M
ꢀ1) was observed when the spec-
Previous study on the photophysical properties of phenazine,
whose structure is shown in Fig. 1, reveals that it is a weak
fluorophore, but it is strongly phosphorescent (41–44). The lowest
trophotometric experiments were conducted at pH 5 4.0, which is
likely within the experimental error of the measurement.
1
The DNA binding constant for DAP, Kb, obtained from the
equilibrium dialysis experiment was calculated from Kb 5 Cb/
[Cf(Stotal ꢀ Cb)], where Cb and Cf are the concentrations of DAP
bound to the DNA and free in solution after the dialysis, re-
spectively, and Stotal represents the total nucleic acid concentration
(25–27,48–51). The total DAP and nucleic acid concentrations
were 10 and 100 lM, respectively, which resulted in Kb 5 3.5 3
103 Mꢀ1. This value is in excellent agreement with that obtained
from the spectrophotometric method. This low binding constant for
DAP with ds-DNA is indicative of a weak interaction, although the
extended p-system of DAP makes the molecule a good candidate
for strong binding through intercalation. Common DNA interca-
lators such as EtBr, proflavin and daunomycin exhibit DNA
energy singlet exited state, S1, of D2h phenazine is B1u (np*). It
has been noted that there is a significant overlap of phenazine’s
3
1B1u (np*) state with its B3u (pp*) state, which is the lowest
energy triplet excited state of the molecule, T1. Because of this
1
3
mixing of the B1u and B3u states, the intersystem crossing rate
from S1 to T1 is much faster than the radiative decay from S1 to the
ground state, S0 (41,42). However, incorporation of substituents
such as amino groups, on the aromatic rings of phenazine, results
in lowering of the np* S1 energy relative to pp* T1, thus increasing
the fluorescence quantum yield and shifting the emission and
absorption maxima to lower energies (42,43). These results are
consistent with the strong fluorescence observed for DAP.
binding constants ;105
M
ꢀ1, which are ;100-fold greater than
DNA binding
that of DAP (51–53).
As shown in Fig. 3a, after addition of ds-DNA to solutions
The binding constant of DAP to ds-DNA was determined using both
equilibrium dialysis and spectrophotometric titrations. To avoid
containing 50 lM DAP (50 mM NaCl, 10 mM phosphorus buffer,

92 Patty K.-L. Fu et al.
Figure 4. EtBr-stained agarose gel (2%) showing the photocleavage of 120
lM pUC19 plasmid by 30 lM DAP after 30 min incubation and 30 min of
irradiation in 10 mM phosphate buffer, pH 5 7.5. a: Lane 1, plasmid only,
dark; Lane 2, plasmid only, irradiated; Lane 3, plasmid þ DAP, dark; Lane
4, plasmid þ DAP, irradiated; Lane 5, plasmid þ EcoR1 restriction enzyme.
b: Lane 1, plasmid only, dark; Lane 2, plasmid only, irradiated; Lanes 3–7,
plasmid þ DAP photolyzed with visible light for 30 min with Lane 3,
pH 5 4.0; Lane 4, pH 5 5.0; Lane 5, pH 5 6.0; Lane 6, pH 5 7.0 and
Lane 7, pH 5 8.0.
Figure 3. a: Relative emission of 50 lM DAP (kexc 5 430 nm, kem 5 590
nm) in 50 mM NaCl, 10 mM phosphorus buffer, pH 5 7.5, as a function of
added CT DNA (5% error in reproducibility). b: Change in the relative
viscosity of 2.5 mM sonicated herring sperm DNA solution with increasing
concentration of EtBr ( ), DAP (r) and Hoechst 33258 (m). A low-salt
n
buffer (25 mM sodium phosphate, pH 7.2) was used.
groove binder Hoechst 33258 does not result in significant changes
in the relative viscosity (56,57), which is typical for minor groove
and electrostatic binders that do not intercalate between the DNA
bases (29,31–33,54–57,60,61). The viscosity data taken together
with the hypochromic shift and luminescence enhancement upon
addition of DNA are in accord with an intercalative binding mode
for DAP. The slope from the EtBr data (;0.91) in Fig. 3b is very
close to that predicted for an ideal intercalator, 1.0 (58). In contrast,
the slope observed for DAP is ;0.57, which can be explained by its
low binding constant to DNA. It is also possible that DAP has other
binding modes to ds-DNA in addition to intercalation, such as
hydrogen bonding through the two amino groups. A slope
significantly lower than 1.0 in the viscosity changes as a function
of additive concentration has been related previously to multiple
binding modes, one of which is intercalation (30,59).
pH 5 7.5), an emission enhancement is observed. Luminescence
enhancement is often observed with emissive intercalators.
Enhanced luminescence can result from p-stacking interaction
with the DNA bases and protection of the intercalator from
aqueous environment. The emission enhancement of DAP upon
binding to DNA is consistent with intercalation (54–59). Although,
a competitive binding study with EtBr can be undertaken to
confirm intercalation (52,53,60,61), the large difference in binding
constant between DAP and EtBr (;5 3 105 Mꢀ1) precludes the use
of this technique for this particular system (51,52). However, the
observed emission enhancement of DAP upon DNA binding is
consistent with protection of the molecule from the surrounding
water, which is typical of molecules that intercalate between the
bases (54,55,58,59). It has been reported that DAP has a more
intense fluorescence in nonpolar solvents, such as acetone and
diethyl ether, than in water, a difference attributed to its use of
water as a source of protons (6,34,42–44).
DNA photocleavage
Changes in relative viscosity provide a reliable method for the
assignment of DNA binding modes by intercalators and groove
binders (56,57). The changes in the relative viscosity of solutions
containing 2.5 mM herring sperm DNA upon addition of increasing
concentrations of DAP are shown in Fig. 3b and parallel those
observed for the intercalator EtBr. In contrast, addition of the minor
The image of the EtBr-stained gel in Fig. 4a shows the DNA
photocleavage of 120 lM pUC19 plasmid by DAP. The control
lanes show that irradiation of the plasmid (pUC19) alone with
visible light for 5 J/cm2 does not result in DNA damage (Lanes 1 and
2). In addition to the supercoiled plasmid (Form I), a small amount
of circular (nicked, Form II) is also found in the control lanes.

Photochemistry and Photobiology, 2005, 81 93
Linearization of the pUC19 supercoiled plasmid with the EcoR1
restriction enzyme results in the exclusive formation of cut, linear
(Form III) ds-DNA, as shown in Lane 5 (Fig. 4a). As shown in Fig.
4a, the addition of 30 lM DAP to 120 lM pUC19 plasmid does not
result in cleavage in the dark (Lane 3); however, irradiation of the
mixture for 5 J/cm2 with visible light (kirr 5 400–700 nm, 30 min)
results in the formation of nicked ds-DNA (Lane 4, Form II).
Because the protonation and deprotonation states of the DAP
have been shown to have a profound effect on the excited-state
properties, the efficiency of DNA cleavage upon irradiation was
measured as a function of pH (Fig. 4b). As shown in Fig. 4b, in the
pH range from 3.0 to 8.0, irradiation of 120 lM pUC19 plasmid in
the presence of 30 lM DAP with visible light results in the
formation of ;28% nicked plasmid at pH 8. A marked decrease in
nicked DNA is evident with the decrease of the pH from 8.0 to 4.0
(Fig. 4b). The results are consistent with greater DNA photo-
cleavage sensitized by DAP in its deprotonated state, where greater
reactivity is observed at pH values of 7 and 8, and a marked
decrease in cleavage is evident with the decrease of the solution pH
from 6 to 4. It is important to note that because of the weak
interaction between DAP and ds-DNA, a 30 min incubation before
irradiation is needed for any DNA photocleavage to occur.
Figure 5. Toxicity tests of DAP. Human skin fibroblasts were incubated in
the dark (cytotoxicity, r) or irradiated (photocytotoxicity, s) with various
concentrations of DAP for 30 min. Cell cultures were irradiated with visible
light (400–700 nm) with the light intensity of 5 J/cm2. Interpolation of the
data points was used to determine the concentration of DAP required to
achieve 50% cell death, LC50. Each point represents the mean of four
determinations and 65% error.
form (pH 5 8), and the emission intensity decreases by an order of
magnitude at pH 5 3. Enhancement of the emission and the
increase in viscosity upon binding of DAP to ds-DNA establishes
that this molecule intercalates between the bases in DNA with a
DAP exhibits DNA photophysical and photobiological charac-
teristics similar to many fluorescent DNA dyes regardless of its
weak binding constant with ds-DNA. It has been demonstrated that
many photoactive DNA dyes are able to intercalate between
nucleobases and nick plasmid DNA in the presence of molecular
oxygen and visible light (62–65). The search for molecules with
desirable photophysical properties, which can function as efficient
artificial photonucleases and biomolecular probes, has resulted in
the discovery of a large number of DNA intercalators that can
cleave ds-DNA via photochemical pathways (66–72). Even though
DAP has proven to possess interesting photosensitizing character-
istics upon irradiation with visible light, it is important to determine
whether it has any functional role in biological systems.
binding constant of 3.5 3 103
M
ꢀ1. However, the changes in
relative viscosity point at multiple binding modes, one of which is
intercalation. Furthermore, DAP is able to produce single-strand
nicks in plasmid DNA upon irradiation with visible light. The
cytotoxicity of DAP toward human skin fibroblasts (Hs-27) was
evaluated both in the dark and upon exposure of the cell cultures
with various concentrations of DAP to visible light. It was observed
that DAP increases the sensitivity of human skin fibroblasts toward
visible light by a factor of 13, resulting in LC50 values of 172 and 13
lM in the dark and upon irradiation, respectively. The excited-state
intermediates responsible for the photocytotoxicity of DAP remain
unknown; however, preliminary experiments show that sodium
azide, a known scavenger of singlet oxygen, results in the complete
inhibition of the DNA photocleavage by DAP (supplementary
material). It is well known that intercalator aromatic amines usually
cleave DNA through the generation of reactive oxygen species
(67,70–73). The potential presence of DAP as a contaminant in
consumer products and with subsequent topical human exposure on
sun-exposed skin requires further investigation of its harmful
effects and photodamage to human cells.
Photocytotoxicity
Irradiation of human skin fibroblasts (Hs-27) exposed to 5–25 lM
DAP with 400–700 nm light for 30 min (5 J/cm2) results in the
death of 25–83% of the cells, respectively, as shown in Fig. 5. In
contrast, .95% cell survival was observed in solutions containing
similar DAP concentrations that were not irradiated with visible
light (Fig. 5). Without irradiation, the survival rate of Hs-27 at
concentrations of DAP up to 125 lM was ;65%.
There is a striking difference in potency between the photo-
cytotoxicity and cytotoxicity of DAP toward Hs-27 cells. The
concentration of DAP, which results in 50% cell death, the lethal
concentration 50 (LC50), for cultures in the dark and exposed to
visible light was found to be 172 and 13 lM, respectively. These
results clearly show that in addition to the toxicity reported
previously for DAP in the dark; the dye is a potent photosensitizer
whose cytotoxicity increases by a factor of 13 upon exposure to
visible light. Therefore, cells simultaneously exposed to DAP and
visible light, the major component of natural sunlight, may be
subject to significant cellular damage.
Acknowledgements—C.T. thanks the National Institutes of Health (RO1
GM64040-01) for their generous support, and P.K.F. thanks the Office of
Cosmetic and Colors, Center for Food and Applied Nutrition, FDA for
their support of this study.
REFERENCES
1. Watanabe, T., T. Hirayama and S. Fukui (1990) Mutagenicity of
commercial hair dyes and detection of 2,7-diaminophenazine. Mutat.
Res. 244, 303–308.
2. Wagner, E. D., A. Cebulska-Wasilewska, S. Connolly and M. J. Plewa
(1996) Mutagenic analysis of 2,3-diaminophenazine and 2-amino-3-
hydroxyphenazine in Salmonella strains expressing different levels of
o-acetyltransferase with and without plant and mammalian activation.
Mutat. Res. 372, 65–74.
CONCLUSIONS
The ground-state absorption and excited-state emission of DAP are
pH dependent. The molecule fluoresces strongly in its deprotonated

94 Patty K.-L. Fu et al.
3. Køppe, B. (1992) Liquid chromatographic determination of 2-hydroxy-
3- aminophenazine and 2,3-diaminophenazine as impurities in pesticide
formulations containing benomyl or carbendazim. J. Chromatogr. A,
623(1), 168–173.
4. Sarrif, A. M., G. T. Arce, D. F. Krahn, R. M. O’Neil and V. L.
Reynolds (1994) Evaluation of carbendazim for gene mutations in the
Salmonella/Ames plate-incorporation assay: the role of aminophenazine
impurities. Mutat. Res./Gene. Tox. 321, 43–56.
26. Ren, J. and J. B. Chaires (1999) Sequence and structural selectivity of
nucleic acid binding ligands. Biochemistry 38, 16067–16075.
27. Chaires, J. B., J. Ren, M. Henary, O. Zegrocka, G. R. Bishop and
L. Strekowski (2003) Triplex selective 2-(2-naphthyl)quinoline com-
pounds: origins of affinity and new design principles. J. Am. Chem. Soc.
125, 7272–7283.
28. Schmechel, D. E. and D. M. Crothers (1971) Kinetic and hydrodynamic
studies of the complex of proflavine with poly A-poly U. Biopolymers
10, 465–480.
29. Wolfe A., G. H. Shimer Jr. and T. Meehan (1987) Polycyclic aromatic
hydrocarbons physically intercalate into duplex regions of denatured
DNA. Biochemistry 26, 6392–6396.
30. Luedtke, N. W., J. S. Hwang, E. Nava, D. Gut, M. Kol and Y. Tor
(2003) The DNA and RNA specificity of eilatin Ru(II) complexes as
compared to eilatin and ethidium bromide. Nucleic Acid Res. 31,
5732–5740.
31. Satyanarayana, S., J. C. Dabrowiak and J. B. Chaires (1992) Neither D-
nor Ã-tris (phenanthroline) ruthenium binds to DNA by classical
intercalation. Biochemistry 31, 9319–9324.
32. Satyanarayana, S., J. C. Dabrowiak and J. B. Chaires (1993) Tris
(phenanthroline) ruthenium (II) enantiomer interaction with DNA:
mode and specificity of binding. Biochemistry 32, 2573–2584.
33. Scaria, P. V. and R. H. Shafer (1991) Binding of ethidium bromide to
a DNA triple helix. J. Biol. Chem. 266, 5417–5423.
34. Brown, K. C., J. F. Corbett and N. P. Loveless (1979) Spectrophoto-
metric studies on the protonation of hydroxyl and aminophenazines in
aqueous solution. Spectrochim. Acta Part A 35, 421–423.
35. Pileni, M.-P. and M. Gratzel (1980) Light induced redox reaction
of proflavin in aqueous and micellar solution. J. Phys. Chem. 84,
2402–2406.
5. Sun, W., K. Jiao and S. S. Zhang (2001) Electrochemical ELISA for the
detection of cucumber mosaic virus using o-phenylenediamine as
substrate. Talanta 55, 1211–1218.
6. Mekler, V. M. and S. M. Bystryak (1992) Application of o-
phenylenediamine as fluorogenic substance in peroxidase-mediated
enzyme-linked immunosorbent assay. Anal. Chim. Acta 264, 359–363.
7. Hamilton, T. M., A. A. Dobie-Galuska and S. M. Wietstock (1999) The
o-phenylenediamine horseradish peroxidase system: enzyme kinetics in
the gereral chemistry laboratory. J. Chem. Edu. 76, 642–647.
8. Rosso, N. D., B. Szpoganicz and A. E. Martell (1999) Catalytic
oxidation of 1,2-diaminobenzene by the dinuclear cobalt(II)-obisdien
dioxygen complex. Inorg. Chim. Acta 287, 193–198.
9. Tanner, A., L. Bowater, A. S. Fairhurst and S. Bornemann (2001)
Oxalate decarboxylase requires manganese and dioxygen for activity.
J. Biol. Chem. 276, 43627–43634.
10. Zuyun, H., H. Huang, R. Cai and Y. Zeng (1998) Organic solvent
enhanced spectrofluorimetric method for determination of lactase
activity. Anal. Chim. Acta 374, 99–103.
11. Zhang, K., R. Cai, D. Chen and L. Mao (2000) Determination of
hemoglobin based on its enzymatic activity for the oxidation of o-
phenylenediamine with hydrogen peroxide. Anal. Chim. Acta 413,
109–113.
12. Watanabe, T., T. Hirayama and S. Fukui (1989) Phenazine derivatives
as the mutagenic reaction product from o- or m-phenylenediamine
derivatives with hydrogen peroxide. Mutat. Res. 227, 135–145.
13. Cebulska-Wasilewska, A., D. Nowak, W. Niedzwiedz, E. D. Wagner
and M. J. Plewa (1999) Studies on the genotoxic effects of amino-
phenazines using two cellular models and three different methods.
Mutat. Res. 446, 57–65.
14. Cebulska-Wasilewska, A., D. Nowak, D. Niedzwiedz and D. Anderson
(1998) Correlation between DNA and cytogenetic damage induced after
chemical treatment and radiation. Mutat. Res. 421, 83–91.
15. Watanabe, T., T. Kasai, M. Arima, K. Okumura, N. Kawabe and T.
Hirayama (1996) Genotoxicity in vivo of phenazine and amino-
phenazines assayed in the wing spot test and the DNA-repair test
Drosophila melanogaster. Mutat. Res. 369, 75–80.
16. Thomas, K. A. and W. B. Euler (2001) An electrochemical
spectroscopic, and theoretical study of poly(2,3-diaminophenazine).
J. Electroanal. Chem. 501, 235–240.
17. Mekler, V. M. and S. M. Bystryak (1992) Autosensitized oxidation of
o-phenylenediamine in an aqueous buffer solution. J. Photochem.
Photobiol. A: Chem. 65, 391–397.
36. Gesan, V. and R. Ramaraj (2001) Spectral properties of proflavin in
zeolite-L and zeolite-Y. J. Lumin. 92, 167–173.
37. Gabor, G., C. Suneet and D. R. Walt (1995) Sensitivity enhancement of
fluorescent pH indicators using pH-dependent energy transfer. Anal.
Chim. Acta 313, 131–137.
38. Teuber, M., M. Rogner and S. Berry (2001) Fluorescent probes for
non-invasive bioenergetic studies of whole cyanobacterial cells.
Biochim. Biophys. Acta 1506, 31–46.
39. Gangola, P., N. B. Joshi and D. D. Pant (1979) Edge excitation red-shift
and excited-state proton association in acridine. Chem. Phys. Lett. 60,
329–331.
40. Sharma, V. K., P. D. Sahare, R. C. Rastogi, S. K. Ghoshal and D.
Mohan (2003) Excited state characteristics of acridine dyes; acriflavin
and acridine orange. Spectrochim. Acta Part A 59, 1799–1804.
41. Choudhury, S. D. and S. Basu (2003) Singlet state exciplet formation of
phenazine with some aromatic amines. Chem. Phys. Lett. 373, 67–71.
42. del Barrio, J. I., J. R. Rebato and F. M. G. Tablas (1989) Basicity of
phenazine in the first triplet state. J. Phys. Chem. 93, 6836–6837.
43. Chalvet, O., H. H. Jaffe and J. C. Rayez (1977) Acid-base equilibria of
electronically excited phenazine. Photochem. Photobiol. 26, 353–356.
44. Aaron, J. J. and M. Maafi (1995) Quantitative treatment of the solvent
effects on the electronic absorption and fluorescence spectra of
acridines and phenazines. The ground and first excited singlet-state
dipole moments. Spectrochim. Acta Part A 51, 603–615.
45. Dietrich, B., R. Ghahary and H. Pasch (1996) Triazine-based polymers.
4. MALDI-MS of triazine-based polyamines. Polymer 37, 777–783.
46. Poulios, I., D. Sazou and G. Kokkinidis (1988) Photoelectrochemical
behavior of electrogenerated poly(2,3-diaminophenazine) on platinum
in aqueous media. Synth. Metals 26, 339–347.
47. Crank, G. and M. I. H. Makin (1989) Photochemical conversion of o-
phenylenediamine and o-aminophenol into heterocyclics. J. Heterocy-
clic Chem. 26, 1163–1166.
48. Fu, K.-L. P. and C. Turro (2001) Transcription inhibition and DNA
photocleavage by metal complexes. Doctoral thesis, The Ohio State
University, Columbus, OH.
49. Fu, K.-L. P., P. M. Bradley and C. Turro (2003) Stabilization of duplex
DNA structure and suppression of transcription in vitro by bis(quinone
diimine) complexes of rhodium(III) and ruthenium(II). Inorg. Chem.
42, 878–884.
50. Sorasaenee, K., P. K.-L. Fu, A. M. Angeles-Boza, K. R. Dunbar and
C. Turro (2003) Inhibition of transcription in vitro by anticancer active
dirhodium(II) complexes. Inorg. Chem. 42, 1267–1271.
18. Bystryak, S. M. and V. M. Mekler (1992) Photochemical amplification
for horseradish peroxidase-mediated immunosorbent assay. Anal. Bio-
chem. 202, 309–393.
19. Mekler, V. M., O. V. Belonogova and V. I. Nikulin (1995) Photochemical
mimetics of chromogenic catalymetric reactions of arylamines in water
solutions. J. Photochem. Photobiol. A: Chem. 87, 243–247.
20. Doyle, R. P., P. E. Kruger, P. R. Mackie and M. Nieuwenhuyzen (2001)
Phenazine-2,3-diamine. Acta Crystallogr. C. 57, 104–105.
21. Zang, K., L. Y. Mai and R. X. Cai (2000) Stopped-flow spectropho-
tometric determination of hydrogen peroxide with hemoglobin as
catalyst. Talanta 51, 179–186.
22. Yuan, H., R. Cai and Z. Pan (2003) Catalitical kinetic determination of
hydrogen peroxide using iron-tetrasulfonatophthalocyanine (with r-CD)
as a mimetic enzyme of peroxidase. Anal. Lett. 36, 277–286.
23. Jang, D.-H., Y.-S. Yoo and S. M. Oh (1995) Electropolymerization
mechanism for poly(o-phenylenediamine) (PPD) and its electrocatalytic
behavior for O2 reduction. Bull. Korean Chem. Soc. 16, 392–397.
24. Sambrook, J. and D. W. Russel (2001) Molecular Cloning, A
Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold
Spring Harbor, New York.
25. Baruah, H. and U. Bierbach (2003) Unusual intercalation of acridin-9-
ylthiourea into the 59-GA/TC DNA base step from the minor groove:
implications for the covalent DNA adduct profile of a novel platinum-
intercalator conjugate. Nucleic Acids Res. 31, 4138–4146.
51. Billadeau, M. A. and H. Morrison (1995) Photoaquation of cis-
Dichlorobis-(1,10-phenanthroline) chromium (III) and the photochemical

Photochemistry and Photobiology, 2005, 81 95
and thermal reactions of this complex with native calf-thymus DNA.
J. Inorg. Biochem. 57, 249–270.
applying time-resolved fluorescence measurement of dye-DNA com-
plexes. Chem. Commun. 8, 689–690.
64. Cosa, G., K.-S. Focsaneanu, J. R. N. McLean, J. P. McNamee and J. C.
Scaiano (2002) Photophysical properties of fluorescent DNA-dyes
bound to single- and double-stranded DNA in aqueous buffered
solution. Photochem. Photobiol. 73, 585–619.
65. Birnboim H. C. and J. J. Jevcak (1981) Fluorimetric method for rapid
detection of DNA strand breaks in human white blood cells produced
by low doses of radiation. Cancer Res. 41, 1889–1892.
66. Rogers, K. R., A. Apostol, S. J. Madsen and C. W. Spencer (1999)
Detection of low dose radiation induced DNA damage using
temperature differential fluorescence assay. Anal. Chem. 71, 4423–
4426.
67. Steullet, V., S. Edwards-Bennett and D. W. Dixon (1999) Cleavage of
abasic sites in DNA by intercalator-amines. Bioorg. Med. Chem. 7,
2531–2540.
68. Fernandez-Saiz, M., P. T. Henderson, W. D. Wilson and G. B. Schuster
(1999) Selective photocleavage of DNA and RNA by anthraquinone
derivatives: targeting the single-strand region of hairpin structures.
Photochem. Photobiol. 70, 847–852.
69. Reynisson, J., G. B. Schuster, S. B Howerton, L. D. Williams, R. N.
Barnett, C. L. Cleveland, U. Landman, N. Harrit and J. B. Chaires
(2003) Intercalation of trioxatriangulenium ion in DNA: binding,
electron transfer, x-ray crystallography, and electronic structure. J. Am.
Chem. Soc. 26, 2072–2083.
70. Turro, C., D. B. Hall, W. Chen, H. Zuilholf, J. K. Barton and N. J.
Turro (1998) Photoreactivity of phenanthrenequinone diimine com-
plexes of rhodium and correlations with DNA photocleavage and
photooxidation. J. Phys. Chem. A 102, 5708–5715.
52. Rosu, F., E. D. Pauw, L. Guittat, P. Alberti, L. Lacroix, P. Mailliet, J.-F.
Riou and J.-L. Mergny (2003) Selective interaction of ethidium
derivatives with quadruplexes: an equilibrium dialysis and electrospray
ionization mass spectrometry analysis. Biochemistry 42, 10361–10371.
53. Qu, X. and J. B. Chaires (2001) Hydration changes for DNA
intercalation reactions. J. Am. Chem. Soc. 123, 1–7.
54. Vaidyanathan, V. G. and B. U. Nair (2003) Synthesis, characterization
and binding studies of chromium (III) complex containing an
intercalating ligand with DNA. J. Inorg. Biochem. 95, 334–342.
55. Hiort, C., P. Lincoln and B. Norden (1993) DNA-binding of D- and Ã-
[Ru(phen)2DPPZ]2þ. J. Am. Chem. Soc. 115, 3448–3454.
56. Suh, D., Y.-K. Oh and J. B. Chaires (2001) Determining the binding mode
of DNA sequence specific compounds. Process Biochem. 37, 521–525.
57. Haq, I., P. Lincoln, D. Suh, B. Norden, B. Z. Chowdhry and J. B. Chaires
(1995) Interaction of DELTA—and LAMBDA [Ru(phen)2DPPZ]2þ
with DNA: a calorimetric and equilibrium binding study. J. Am. Chem.
Soc. 117, 4788–4796.
58. Cohen, G. and H. Eisenbrg (1969) Viscosity and sedimentation study of
sonicated DNA-proflavin complexes. Biopolymers 8, 45–55.
59. Jin, E., V. Katritch, W. K. Olson, M. Kharatisvil, R. Abagyan and D. S.
Pilch (2000) Aminoglycoside binding in the major groove of duplex
RNA: the thermodynamic and the electrostatic forces that govern
recognition. J. Mol. Biol. 298, 95–110.
60. Tang, T. C. and H. J. Huang (1999) Electrochemical studies of
the intercalation of ethidium bromide to DNA. Electroanalysis. 11,
1185–1190.
61. Xie, H. P., J. H. Jiang, X. Chu, H. Cui, H. L. Wu, G. L. Shen and R. Q.
Yu (2002) Competitive interaction of the antitumor drug daunorubicin
and the fluorescence probe ethidium bromide with DNA as studied by
resolving trilinear fluorescence data: the use of PARAFAC and its
modification. Anal. Bioanal. Chem. 373, 159–162.
62. Jennette, K. W., S. J. Lippard, G. A. Vassiliades and W. R. Bauer
(1774) Metallointercalation reagents 2-hydroxyethanethiolato(2,29,29-
terpyridine)-platinum(II) monocation binds strongly to DNA by
intercalation. Proc. Natl. Acad. Sci. USA 71, 2829–2843.
71. Delaney, S., M. Pascaly, P. K. Bhattacharya, K. Han and J. K. Barton
(2002) Oxidative damage by ruthenium complexes containing the
dipyridophenazine ligand or its derivatives: a focus on intercalation.
Inorg. Chem. 41, 1966–1974.
72. Copeland, K. D., A. M. Lueras, E. D. Stemp and J. K. Barton (2002)
DNA cross-linking with metallointercalator-peptide conjugates. Bio-
chemistry 22, 12785–12797.
73. Kornhauser, A., W. G. Wamer and L. A. Lambert (2002) Cellular and
molecular events following ultraviolet irradiation of skin. In Dermato-
toxicology (Edited by F. N. Marzulli and H. I. Maibach), pp. 189–230.
Taylor & Francis, Washington, DC.
63. Cosa, G., K.-S. Focsaneanu, J. R. N. McLean and J. C. Scaiano (2000)
Direct determination of single to double stranded DNA in solution