nihms97873.pdf

DNA Mismatch Binding and Antiproliferative Activity of Rhodium

Metalloinsertors

Russell J. Ernst

Hang Song

, and

Jacqueline K. Barton

Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena,

California 91125

Abstract

Deficiencies in mismatch repair (MMR) are associated with carcinogenesis. Rhodium

metalloinsertors bind to DNA base mismatches with high specificity and inhibit cellular proliferation

preferentially in MMR-deficient cells versus MMR-proficient cells. A family of chrysenequinone

diimine complexes of rhodium with varying ancillary ligands that serve as DNA metalloinsertors

has been synthesized, and both DNA mismatch binding affinities and antiproliferative activities

against the human colorectal carcinoma cell lines HCT116N and HCT116O, an isogenic model

system for MMR deficiency, have been determined. DNA photocleavage experiments reveal that all

complexes bind to the mismatch sites with high specificities; DNA binding affinities to

oligonucleotides containing single base CA and CC mismatches, obtained through photocleavage

titration or competition, vary from 10

to 10

−

for the series of complexes. Significantly, binding

affinities are found to be inversely related to ancillary ligand size and directly related to differential

inhibition of the HCT116 cell lines. The observed trend in binding affinity is consistent with the

metalloinsertion mode where the complex binds from the minor groove with ejection of mismatched

base pairs. The correlation between binding affinity and targeting of the MMR-deficient cell line

suggests that rhodium metalloinsertors exert their selective biological effects on MMR-deficient cells

through mismatch binding

in vivo

Introduction

The mismatch repair (MMR) pathway corrects single base errors and insertion/deletion loops

that arise during DNA synthesis, increasing the fidelity of DNA replication by a factor of 50–

1000.

If uncorrected, mismatches are converted to mutations in subsequent cycles of DNA

replication, and cells with MMR deficiencies, not surprisingly, exhibit elevated mutation rates.

–

Germline mutations in

hMLH1

hMSH2

, essential genes for MMR in humans,

dramatically increase the risk of developing hereditary nonpolyposis colon cancer (HNPCC),

the most common type of inherited colon cancer.

HNPCC is marked by early onset and the

presence of cancers in several other tissue types.

Roughly 15% of sporadic colorectal cancer

cases have also been linked to MMR deficiency.

Epigenetic silencing of the MMR genes has

been identified as the cause of MMR deficiency in these cases.

In addition to colorectal cancer,

mismatch repair deficiencies have been found in approximately 16% of solid tumors of all

tissue types.

E-mail: jkbarton@caltech.edu.

Supporting Information Available:

Figures S1 and S2, showing inhibitory effects of

rac

-Rh(HDPA)

chrysi

and

rac

-Rh

(phen)

chrysi

as a function of incubation time on cellular proliferation. This material is available free of charge via the Internet at

http://pubs.acs.org.

NIH Public Access

Author Manuscript

J Am Chem Soc

. Author manuscript; available in PMC 2010 February 18.

Published in final edited form as:

J Am Chem Soc

. 2009 February 18; 131(6): 2359–2366. doi:10.1021/ja8081044.

NIH-PA Author Manuscript

Importantly, MMR deficiency confers resistance or tolerance to many of the anti-cancer agents

currently in clinical use.

Alkylation by the commonly used chemotherapeutic agents

methyl-

-nitrosourea (MNU) and

-methyl-

′

-nitro-

-nitrosoguanidine (MNNG) at the O6

position of guanine nucleotides triggers an apoptotic response after recognition of O6-meG:C

and O6-meG:T base pairs by the MMR pathway, while MMR-deficient cells tolerate this DNA

methylation.

–

Failure to recognize DNA adducts is also involved in the resistance of MMR-

deficient cells to the platinum compounds cisplatin and carboplatin.

–

The incorporation of

anti-metabolites such as 5-fluorouracil and 6-thioguanine into DNA triggers cell cycle arrest

and apoptosis through the MMR pathway, and consequently MMR-deficient cells are resistant

to these agents as well.

Other studies have shown low-level resistance to the type I

topoisomerase poisons camptothecin and topotecan in

hMLH1

deficient lines and to the type

II topoisomerase poisons doxorubicin, epirubicin, and mitoxantrone in

hMLH1

hMSH2

deficient lines.

It has also been hypothesized that treatment regimens with agents such as

cisplatin might enrich tumors for MMR-deficient cells,

and it has been shown that a

substantial portion of secondary, or therapy-related, leukemias show signs of MMR deficiency.

Collectively, these results show the broad involvement of MMR in mediating drug

response, the effects of MMR deficiency on this response, and the need to develop therapeutic

agents that specifically target MMR-deficient cells.

Our laboratory has previously developed bulky rhodium complexes that target DNA

mismatches

in vitro

–

These octahedral complexes contain an expansive tetracyclic

aromatic ligand that can be accommodated preferentially by DNA at a thermodynamically

destabilized mismatch site. The first-generation compound, Rh(bpy)

chrysi

(chrysi = 5,6-

chrysenequinone diimine), binds 80% of DNA mismatches with typical binding constants of

−

and remarkable specificity for mismatched DNA; in a 2.6 kb DNA fragment DNA

photocleavage reveals specific targeting of the mismatch.

Subsequent work led to the

incorporation of nitrogen atoms into the intercalating ligand and a 50-fold increase in binding

affinity for the second-generation compound, Rh(bpy)

phzi

(phzi = benzo[

]-phenazine-5,6-

quinonediimine).

A high-resolution crystal structure of Rh(bpy)

chrysi

bound to single

AC mismatches within a DNA oligonucleotide duplex reveals a distinctive binding mode at

the mismatched site.

We had previously determined that tris(chelate) complexes of Rh with

a planar aromatic ligand bind to well-matched DNA by partial intercalation of the planar ligand

from the major groove side into the base pair stack.

However, binding to the mismatched

site involves instead insertion of the expansive ligand into the DNA duplex from the minor

groove side at the mismatched site with ejection of the mismatched bases out of the DNA stack;

the inserted ligand stacks fully with adjacent base pairs. NMR studies of Rh(bpy)

chrysi

bound to an oligonucleotide containing a CC mismatch confirm this metalloinsertion mode for

the complex at mismatched sites in solution.

The

in vivo

effects of Rh(bpy)

chrysi

and Rh(bpy)

phzi

have been characterized in the

isogenic cell lines HCT116N and HCT116O.

The HCT116 cell line is a colorectal carcinoma

line deficient in the

hMLH1

gene. Two derivative cell lines, HCT116N and HCT116O, have

been made through transfection of human chromosome 3 (ch3) and human chromosome 2

(ch2), respectively. The presence of a functional copy of ch3 restores MMR proficiency in the

HCT116N line, while the HCT116O line transfected with ch2 remains MMR deficient.

The

mismatch recognition compounds developed within our laboratory were shown to selectively

inhibit the proliferation of the repair-deficient HCT116O line.

Recent work within our laboratory on luminescent ruthenium complexes has also shown that

these tris(chelate) complexes are taken up inside cells through passive diffusion facilitated by

the membrane potential.

Variations in ancillary ligands have dramatic effects on cellular

uptake, with increased lipophilicity facilitating uptake. Uptake can also be increased through

functionalization with a nuclear localizing peptide.

Ernst et al.

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NIH-PA Author Manuscript

Here we examine the effects of ancillary ligand variation in the Rh(L)

chrysi

family on the

ability of these complexes to target DNA mismatches

in vitro

and

in vivo

. Importantly, we

establish that the differential inhibition of cellular proliferation in MMR-deficient cells is

correlated with mismatch binding affinity.

Experimental Procedures

Materials

RhCl

was purchased from Pressure Chemical, Inc. (Pittsburgh, PA). [Rh(NH

)

Cl]Cl

was

obtained from Strem Chemical, Inc. (Newburyport, MA). 2,2

′

-Dipyridylamine (HDPA), 4,7-

diphenyl-1,10-phenanthroline (DIP), and Sephadex ion-exchange resin were obtained from

Sigma-Aldrich (St. Louis, MO). Sep-Pak C

solid-phase extraction cartridges were purchased

from Waters Chemical Co. (Milford, MA). Phosphoramidites were purchased from Glen

Research (Sterling, VA). Media and supplements were purchased from Invitrogen (Carlsbad,

CA). BrdU, antibodies, buffers, and peroxidase substrate were purchased in kit format from

Roche Molecular Biochemicals (Mannheim, Germany). All commercial materials were used

as received.

Oligonucleotide Synthesis

Oligonucleotides were synthesized on an Applied Biosystems 3400 DNA synthesizer using

standard phosphoramidite chemistry. DNA was synthesized with a 5

′

-dimethoxytrityl (DMT)

protecting group. The oligonucleotides were cleaved from the beads by reaction with

concentrated ammonium hydroxide at 60 °C overnight. The resulting free oligonucleotides

were purified by HPLC using a C

reverse-phase column (Varian, Inc.) on a Hewlett-Packard

1100 HPLC. The DMT group was removed by reaction with 80% acetic acid for 15 min at

room temperature. The DMT-free oligonucleotides were precipitated with absolute ethanol

and purified again by HPLC. Positive identification of the oligonucleotides and their purity

were confirmed by MALDI-TOF mass spectrometry. Quantification was performed on a

Beckman DU 7400 spectrophotometer using the extinction coefficients at 260 nm (

260

)

estimated for single-stranded DNA.

Synthesis and Characterization of Metal Complexes

Chrysene-5,6-dione, [Rh(bpy)

chrysi]Cl

, and [Rh(phen)

chrysi]Cl

were prepared according

to previously reported procedures.

[Rh(NH

)

chrysi]OTf

[Rh(NH

)

]OTf

was prepared as described by Sargeson.

[Rh(NH

)

]OTf

was reacted

with a limiting amount of chrysene quinone in a 3:1 acetonitrile:water mixture with excess

sodium hydroxide as a catalyst to form [Rh(NH

)

chrysi]OTf

. Acetonitrile was removed

vacuo

, followed by filtration to remove unreacted chrysenequinone. The product was separated

from unreacted [Rh(NH

)

]OTf

by solid-phase extraction on a C

cartridge and eluted with

1:1:0.001 acetonitrile:water:TFA.

H NMR (DMSO-

, 50 °C, 300 MHz):

13.30 (s), 12.32

(s), 8.876 (t, 1H, 7.7 Hz), 8.787 (d, 1H, 7.9 Hz), 8.57–8.51 (m, 2H), 8.358 (dd, 1H, 8.9 Hz, 4.6

Hz), 8.145 (d, 1H, 7.7 Hz), 7.85–7.70 (m, 4H), 4.73–4.54 (broad m, 6H), 3.862 (s, 3H) ppm.

UV/vis (H

O, pH 5): 263 nm (60 900 M

−

), 283 nm (38 100 M

−

), 326 nm (12 600

−

), 413 nm (12 000 M

−

). MALDI-MS (cation): 425

−

) obsd, 427

calcd.

rac

-[Rh(HDPA)

(chrysi)]OTf

[Rh(NH

)

(chrysi)]OTf

(15 mg, 0.02 mmol) was reacted with HDPA (20 mg, 0.12 mmol,

excess) in 20 mL of ethanol and 20 mL of water. The dark red solution was heated under reflux

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NIH-PA Author Manuscript

for 16 h. The reaction mixture turned reddish brown upon heating. Ethanol was removed under

vacuum, and the resulting solution was filtered to remove any residue. The filtrate was

concentrated on a Sep-Pak C

cartridge, eluting with 1:1:0.01 acetonitrile:water:TFA,

lyophilized, and purified on an alumina column, eluting with 2% methanol in dichloromethane.

The fractions were collected and dried under vacuum to give an orange-brown solid (8 mg,

39%).

H NMR (DMSO-

, 300 MHz):

12.84 (s, 1H), 12.34 (s, 1H), 11.78 (s, 1H), 10.32 (d,

1H, 8.7 Hz), 8.63 (d, 1H, 6.9 Hz), 8.40 (d, 1H, 8.4 Hz), 8.31 (d, 1H, 9.3 Hz), 8.14 (m, 2H),

8.07 (d, 1H, 8.7 Hz), 8.04 (d, 1H, 5.4 Hz), 7.94 (m, 4H), 7.77 (m, 5H), 7.58 (m, 2H), 7.48 (d,

1H, 8.1 Hz), 7.41 (d, 1H, 8.4 Hz), 7.32 (s, 1H), 7.14 (m, 2H), 7.04 (t, 1H, 6.8 Hz), 6.98 (t, 1H,

6.9 Hz), 6.81 (t, 1H, 6.5 Hz) ppm. UV/vis (H

O, pH 5): 287 nm (42 200 M

−

), 321 nm

(23 000 M

−

), 442 nm (8800 M

−

). ESI-MS (cation): 699.2

−

), 350.1

−

) obsd, 699.2

−

) calcd.

rac

-[Rh(DIP)

(NH

)

]OTf

RhCl

and 2 equiv of DIP were combined in 1:1 ethanol:water and refluxed overnight. The

solvent was removed

in vacuo

, and the product was recrystallized by dissolving in acetonitrile

at 60 °C and cooling to

−

20 °C. The precipitate was collected by filtration, washed in diethyl

ether, and dissolved in neat triflic acid. The solution was again cooled and added dropwise to

OH at

−

20 °C. The pale white precipitate was collected by filtration and washed with a

small amount of water to give [Rh(DIP)

(NH

)

]OTf

rac

-[Rh(DIP)

chrysi]Cl

rac

-[Rh(DIP)

(NH

)

]OTf

was combined with a 10% excess of 5,6-chrysenequinone and a

catalytic amount of NaOH in acetonitrile and stirred at room temperature overnight. The

condensation was terminated by addition of a stoichiometric amount of HCl. The solvent was

removed

in vacuo

, and the product was purified by alumina column chromatography. Unbound

chrysi ligand eluted first with ethyl acetate, and the purified product then eluted with

acetonitrile. Finally, the compound was dissolved in 3:2 MeCN:H

O, and the triflate counterion

was exchanged for chloride ion with Sephadex QAE-125 ion-exchange resin. UV/vis (H

pH 5): 290 nm (104 000 M

−

), 335 nm (43 900 M

−

), 373 nm (22 300 M

−

). ESI-MS (cation): 1020.9

−

), 511.0

−

) obsd, 1023

−

) calcd.

Photocleavage Titrations

The oligonucleotide was

P-labeled at the 5

′

-end by incubating DNA with

P-ATP and

polynucleotide kinase (PNK) at 37 °C for 2 h, followed by purification using gel

electrophoresis. A small amount of the labeled DNA (less than 1% of the total amount of DNA)

was added to 2

M DNA in 100 mM NaCl, 20 mM NaP

, pH 7.1 buffer. The DNA hairpin was

annealed by heating at 90 °C for 10 min and cooling slowly to room temperature over a period

of 2 h. Racemic rhodium complex solutions ranging from nanomolar to micromolar

concentration were made in Milli-Q water. Annealed 2

M DNA (10

L) and 10

L of Rh

solution at each concentration were mixed in a microcentrifuge tube and incubated at 37 °C

for 10 min. A light control (LC), in which the DNA was mixed with 10

L of water and

irradiated, and a dark control (DC), in which the DNA was mixed with the highest concentration

of rhodium complex without irradiation, were also prepared. The samples were left in the heat

block and irradiated on an Oriel (Darmstadt, Germany) 1000-W Hg/Xe solar simulator (340–

440 nm) for 5 min. The irradiated samples were dried and electrophoresed in a 20% denaturing

polyacrylamide gel. The gel was then exposed to a phosphor screen, and the relative amounts

of DNA in each band were quantitated by phosphorimagery (ImageQuant).

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Binding Constant Determination

The fraction of DNA cleaved in each lane on the gel was normalized and plotted against the

log of the concentration of rhodium complex. At least three photocleavage titrations were

carried out for each racemic metal complex. The pooled data were fit to a sigmoidal curve

using OriginPro 6.1. The resulting midpoint value (i.e., the log of [rhodium complex] at the

inflection point of the curve) was converted to units of concentration ([Rh

50%

]). The

dissociation constant was calculated according to

= [Rh

50%

]

−

0.5[DNA], and the binding

constant was defined as

= 1/

. The errors were derived from the errors associated with

the midpoint values. For complexes that did not photocleave DNA, a binding competition

titration was carried out with a constant amount (1

M) of

rac

-Rh(bpy)

(chrysi)

added to

each sample. The binding and dissociation constants of the non-photocleaving complex were

calculated by solving simultaneous equlibiria involving DNA, Rh(bpy)

(chrysi)

, and the

complex in question in Mathematica 6.0.

Cell Culture

HCT116N and HCT116O cells were grown in RPMI medium 1640 supplemented with 10%

FBS, 2 mM

-glutamine, 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, 100 units/

mL penicillin, 100

g/mL streptomycin, and 400

g/mL Geneticin (G418). Cells were grown

in tissue culture flasks and dishes (Corning Costar, Acton, MA) at 37 °C under 5% CO

atmosphere.

Cellular Proliferation ELISA

HCT116N and HCT116O cells were plated in 96-well plates at 2000 cells/well and allowed

24 h to adhere. The cells were then incubated with rhodium complexes for the durations

specified. For incubation less than 72 h, the Rh-containing medium was replaced with fresh

medium, and the cells were grown for the remainder of the 72 h period. Cells were labeled

with BrdU 24 h before analysis. The BrdU incorporation was quantified by antibody assay

according to established procedures.

Cellular proliferation was expressed as the ratio of

the amount of BrdU incorporated by the treated cells to that of the untreated cells.

Results

Binding Affinities for Metal Complexes at Single Base Mismatches

The binding constants of the family of Rh(L)

chrysi

complexes at a CC and AC mismatch

in a 29-mer DNA hairpin with the sequence 5

′

GGCAGG

XATGGCTTTTTGCCAT

CCCTGCC-3

′

(

X = C or A, underline denotes the

mismatch) were measured. The hairpin sequence allows cleavage site determination on either

strand around the DNA mismatch site. By irradiating samples of DNA titrated with varying

concentrations of a rhodium complex, a photocleavage titration curve is obtained from which

the binding constant of the rhodium complex is determined. A typical autoradiogram, taken

after electrophoresis through a denaturing gel, of samples in a photocleavage titration with

rac

-Rh(bpy)

chrysi

at a CC mismatch is shown in Figure 2. The position of the

photocleavage bands indicates that Rh(bpy)

chrysi

cleaves one base neighboring the

mismatch site near the 3

′

-end. For this DNA sequence, we observed cleavage on only one

strand as reported earlier.

This cleavage pattern is found for both

rac

-Rh(phen)

chrysi

and

rac

-Rh(DIP)

chrysi

and holds for the AC mismatch for

rac

-Rh(bpy)

chrysi

and

rac

-Rh

(phen)

chrysi

. No other photocleavage bands are observed, demonstrating the high

specificity of Rh(L)

chrysi

complexes binding to the mismatch. The photocleavage titration

curve is generated from the autoradiogram by quantifying the amount of photocleavage relative

to the total amount of DNA at each Rh concentration. Pooled data from at least three repeats

were fitted to a sigmoidal curve (Figure 2) for determination of the midpoint ([Rh

50%

]) and

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NIH-PA Author Manuscript

the dissociation constant (

). The

value for

rac

-Rh(bpy)

chrysi

at a CC mismatch is

found to be 30 nM. For

rac

-Rh(phen)

chrysi

and

rac

-Rh(DIP)

chrysi

at a CC mismatch,

values of 320 nM and 11

M are found, respectively (corresponding

values are shown

in Table 1). For the AC mismatch with both

rac

-Rh(bpy)

chrysi

and

rac

-Rh

(phen)

chrysi

values are somewhat higher, as we expect given the greater

thermodynamic stability of an AC mismatch versus a CC mismatch.

rac

-Rh(DIP)

chrysi

does not yield any photocleavage up to 100

M; thus, its

value is estimated to be greater

than that.

As with phenanthrenequinone diimine complexes of rhodium containing saturated amine

ligands, Rh(NH

)

chrysi

and

rac

-Rh(HDPA)

chrysi

promote relatively little DNA

cleavage upon irradiation.

As a result, their binding affinities were determined through

binding competition titrations with 1

rac

-Rh(bpy)

chrysi

. On the basis of the binding

constant of Rh(bpy)

chrysi

, the binding constant of Rh(NH

)

chrysi

is calculated by

solving simultaneous equilibria at the inflection point of the photocleavage titration curve.

Through this competitive titration, the binding constant of Rh(NH

)

chrysi

at a CC mismatch

is found to be 1.0 × 10

−

. At an AC mismatch,

of Rh(NH

)

chrysi

is 3.4 × 10

−

. From similar binding competition titrations, the binding constant of

rac

-Rh

(HDPA)

chrysi

is found to be 2.0 × 10

−

at a CC mismatch and 2.6 × 10

−

at an AC

mismatch. It is apparent that the binding affinity correlates inversely with complex size; the

smallest complex, Rh(NH

)

chrysi

, shows the highest affinities for mismatched sites. The

binding constants for the entire series of Rh(L)

chrysi

complexes are summarized in Table

Inhibition of Cellular Proliferation by Enzyme-Linked Immunosorbent Assay (ELISA)

An ELISA for DNA synthesis was used to quantify the effects of the metalloinsertors on the

proliferation of HCT116N cells (MMR-proficient) and HCT116O cells (MMR-deficient).

Both cell lines were incubated with 0–25

M of each member of the Rh(L)

chrysi

family

except Rh(DIP)

chrysi

, which was administered at 0–5

M concentrations due to its greater

uptake characteristics. Incubations were performed for 12, 24, 48, or 72 h. After the 12, 24,

and 48 h incubations, the medium containing Rh was replaced with fresh medium, and the cells

were grown for the remainder of the 72 h period. The extent of cellular proliferation is expressed

as the ratio of BrdU incorporated by the rhodium-treated cells as compared to untreated

controls. Figure 3 shows representative data for Rh(NH

)

chrysi

at various incubation times.

No significant preferential inhibition of the HCT116O cell line is seen at incubation times less

than 24 h, consistent with previous results for Rh(bpy)

chrysi

, with the exception of Rh

(DIP)

chrysi

, which displays a small differential effect at 12 h.

With longer incubation

times, however, Rh(NH

)

chrysi

displays a strong differential effect with preferential

inhibition of the MMR-deficient HCT116O cell line over the MMR-proficient HCT116N cell

line. In particular, 48 h treatment with 10

M Rh(NH

)

chrysi

inhibits the proliferation of

the HCT116O line by 82 ± 2% while exerting little to no effect on the HCT116N line (7 ± 6%

inhibition).

Figure 4 shows the ELISA results for members of the metalloinsertor family as a function of

incubation time. We have shown previously that the

-enantiomer of Rh(bpy)

chrysi

biologically inactive

and that structurally binding to a mismatch site is enantiospecific for

the

-isomer.

For this reason, treatment with the 10

M achiral tetraammine complex was

compared to treatment with 20

M racemic mixtures of the Rh(L)

chrysi

complexes (L =

HDPA, bpy, or phen). The differential effect of rhodium treatment between the cells lines was

quantified by subtracting the normalized percentages of cellular proliferation for each cell line.

Notably, the optimal incubation time for each compound is inversely related to the

hydrophobicity of the ancillary ligands, with

rac

-Rh(phen)

chrysi

exhibiting an optimal

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incubation time of 24 h. This trend also continues with

rac

-Rh(DIP)

chrysi

, which exhibits

differential effects in as little as 12 h at concentrations as low as 2

M (Figure 5). Based on the

early effect at 12 h, the HDPA complex may have different uptake characteristics; the

analogous Ru complex has not yet been examined using flow cytometry. With the exception

of the HDPA complex, this variation in activity with incubation time for the family of

complexes parallels closely results seen earlier for uptake in HeLa cells by Ru(bpy)

dppz

Ru(phen)

dppz

, and Ru(DIP)

dppz

, where the most rapid uptake is apparent with the

lipophilic DIP complex.

Figure 6 summarizes the differential effects on cell proliferation and the incubation time for

the family of complexes. Clear correlations with the binding constants for these complexes are

evident (Table 1). Significantly, the differential effect in inhibiting cell proliferation in MMR-

deficient cells is directly correlated to the binding affinity of the compound for DNA

mismatches. Rh(NH

)

chrysi

(

= 1 × 10

−

at a CC mismatch), for example, shows the

largest differential effect in inhibiting proliferation of MMR-deficient versus -proficient

HCT116 cells after 72 h (79 ± 5%), while Rh(phen)

chrysi

(

= 3.2 × 10

−

at a CC

mismatch) shows a small differential effect (17 ± 7%). The DIP complex is rapidly taken up

by the cells but also shows only a small differential inhibitory effect correlating with its poor

specific binding at the mismatch site.

Discussion

A clear trend emerges when comparing the binding constants of the series of rhodium

complexes to mismatched sites: the DNA mismatch binding affinity increases as the size of

the ancillary ligand decreases. This trend is consistent with what we have learned from the

structural studies, specifically that mismatch binding by insertion via the minor groove is

subject to stringent space constraints. With major groove intercalation, the base rise is increased

and the major groove offers space to accommodate the ancillary ligands. In contrast, with

insertion, there is no increase in base pair rise; the mismatched bases are instead ejected and

replaced by the deeply inserted chrysi ligand. Moreover, the minor groove, small even for

hydrophobic groove binding molecules, offers little space for the ancillary ligands. While little

enantioselectivity is apparent for intercalation of bpy complexes into B-form DNA,

-Rh

(bpy)

chrysi

binds enantiospecifically to single base mismatches.

Thus steric

interactions of the ancillary ligands are seen as an extremely important factor governing the

binding affinity of a metal complex at the mismatch site.

We have previously demonstrated that mismatch binding affinity is correlated with

thermodynamic destabilization over all mismatch identities and sequence contexts.

Here,

we see for all the complexes that binding to the CC mismatch is tighter than binding to the AC

mismatch. This is consistent with our previous observations, since AC is the

thermodynamically more stable mismatch and, in this case, is estimated to stabilize the hairpin

duplex by

∼

0.5 kcal/mol relative to one containing a CC mismatch.

We assume, then, that

the general trend holds for all members of this metalloinsertor family. This stabilization is

translated into a higher dissociation constant (smaller binding affinity) for the entire series of

rhodium complexes. This decrease in binding affinity depends upon the greater energy required

to eject the mismatched bases from the base pair stack, as evident crystallographically and by

NMR.

Nonetheless, for the family of chrysi complexes, the inverse relationship between

the size of the ancillary ligand and the binding affinity still holds, with the smallest complex

Rh(NH

)

chrysi

showing the highest affinity and that of the largest complex, Rh

(DIP)

chrysi

, more than 2 orders of magnitude lower.

Figure 7 compares the crystal structure of

-Rh(bpy)

chrysi

bound to the mismatch site

with models of

-Rh(DIP)

chrysi

and Rh(NH

)

chrysi

similarly bound via the minor

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groove through metalloinsertion. Preserving the DNA conformation from the crystal structure,

we see that

-Rh(DIP)

chrysi

runs into substantial steric hindrance, as its axial phenyl rings

extend up and down into the groove, directly clashing with the bases. However, its equatorial

phenyl rings do not pose any steric problems, as they point away from the DNA. These

observations are supported by the small binding constant measured for Rh(DIP)

chrysi

. Rh

(phen)

chrysi

, intermediate in size, shows binding affinities for the mismatches that are an

order of magnitude lower than those of the bpy derivative but more than an order of magnitude

higher than those of the DIP complex. Rh(HDPA)

chrysi

is slightly larger in size than the

bpy derivative, but the HDPA ligands are more flexible, and there is an opportunity for

hydrogen bonding; as a result, Rh(bpy)

chrysi

and Rh(HDPA)

chrysi

have comparable

affinities for the mismatch. Analogously, the large binding constant of Rh(NH

)

chrysi

can

be mostly attributed to its small size. Here it is reasonable to suggest that the axial ammines

may also hydrogen bond with the neighboring base pairs to form additional stabilizing

interactions. Nonetheless, as evident in Figure 7, the small cone size of the tetraammine

structure clearly facilitates deep insertion within the minor groove site. In fact, the clear inverse

correlation of binding affinity with ancillary ligand size, and the finding that Rh

(DIP)

chrysi

, despite its cumbersome size, is able to bind at all to a mismatch site with some

specificity, corroborate our understanding of the driving force and the dynamics of mismatch

recognition: the

-stacking between the inserted chrysi ligand and the adjacent bases provides

the major stabilizing force for binding, and both the metal complex and DNA distort their

conformations to accommodate each other in the bound state.

Importantly, the DNA mismatch binding affinities of the Rh(L)

chrysi

family correlate well

with the differential biological effects seen between the repair-proficient HCT116N and repair-

deficient HCT116O cell lines. This correlation supports the hypothesis that DNA mismatches

are the target of rhodium metalloinsertors

in vivo

. Because of this correlation, we may attribute

the preferential inhibitory effect on MMR-deficient cells to binding of the complexes to DNA

mismatches. Since the MMR-deficient cells contain more mismatches, the tighter binding

complexes would be expected to display a greater inhibitory effect. It should be noted that

finding any inhibitory effect with these complexes was at first surprising, since they bind DNA

noncovalently and might be expected to be readily displaced. Although the mechanism of

inhibition is not yet fully understood, it is likely that protein recognition of the metal-mismatch

complex, perhaps by RNA polymerase or topoisomerase, may generate a covalent protein–

DNA lesion and contribute to the cellular response.

The differential inhibitory effect seen with Rh(HDPA)

chrysi

cannot be understood simply

on the basis of binding affinities. Despite having essentially the same mismatch binding affinity

as Rh(bpy)

chrysi

, the HDPA complex preferentially inhibits the MMR-deficient cell line

almost as well as Rh(NH

)

chrysi

with long incubation times; with short times of incubation,

the differential inhibitory effect by Rh(HDPA)

chrysi

is greatest. Both the HDPA ligand and

the amine group have the potential to form hydrogen bonds. This hydrogen bonding capability

and flexibility of the ligands might serve to make them more effective inhibitors of any protein–

DNA interactions. Indeed, ruthenium complexes bearing HDPA ligands have been shown to

exhibit DNA binding and cytotoxicity.

Certainly, as with any pharmaceutical design, cellular uptake must also be considered. In the

case of the HDPA complex, based upon the variations in inhibitory effect with incubation time,

the amine ligands may facilitate nuclear uptake. Dppz analogues with the HDPA ligands have

not yet been examined with respect to their uptake characteristics. For the bpy complexes, the

48 h incubation time required for Rh(bpy)

chrysi

to exert its anti-proliferative effect matches

the 48 h requirement observed for Ru(bpy)

dppz

uptake in HeLa cells.

The more

lipophilic Rh(DIP)

chrysi

here is found to exert anti-proliferative effects at much shorter

incubation times and lower concentrations, which also matches the accelerated uptake observed

Ernst et al.

Page 8

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NIH-PA Author Manuscript

for Ru(DIP)

dppz

. Cellular uptake is surely a rate-limiting factor in biological activity of the

rhodium metalloinsertors, yet cellular uptake is not the only challenge: proper subcellular

localization must also be achieved in order for any drug to act upon its target. It has been well

established that lipophilic cations preferentially target the mitochondria, whereas hydrophilic

cations do not.

It may be that Rh(HDPA)

chrysi

and Rh(NH

)

chrysi

lack the

lipophilicity required for mitochondrial accumulation, allowing a greater proportion of these

compounds to reach the nucleus once inside the cell. This difference in intracellular partitioning

could then account for the differential effects of Rh(HDPA)

chrysi

In the development of octahedral rhodium complexes as anti-cancer agents, the choice of

ancillary ligand can be seen as a design tradeoff, with the binding affinity for a DNA mismatch

greatly outweighing uptake properties as the critical factor in the successful targeting of repair-

deficient cells. Beyond their effects on DNA binding and overall cellular uptake, it is highly

likely that the ancillary ligands affect the cellular response in other ways, including the potential

for hydrogen bonding and differences in uptake and intracellular distribution. Here we are

confronted with a tradeoff that may seem inevitable: more hydrophobic ligands facilitate

cellular uptake but impede mismatch binding. Perhaps this tradeoff can be avoided by making

conjugates arranged with functional moieties tethered with consideration of the structure of

the DNA-bound complex associated snugly in the minor groove. Most importantly, these data

support the contention that the cell-specific inhibitory effect we observe depends upon binding

to the DNA mismatch inside the cell. This cell-specific strategy thus represents a promising

direction in the development of small metal complexes that react preferentially in MMR-

deficient cells, those susceptible to cancerous transformation.

Supplementary Material

Refer to Web version on PubMed Central for supplementary material.

Acknowledgments

Financial support for this work from the NIH (GM33309 to J.K.B.) is gratefully acknowledged. The authors also thank

Dr. Jonathan Hart for his synthesis of [Rh(NH

)

chrysi]OTf

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Figure 1.

Rh(L)

chrysi

family of metalloinsertors.

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Figure 2.

Binding affinities determined through DNA photocleavage. The DNA hairpin sequence is 5

′

GGCAGG

XATGGCTTTTTGCCAT

CCCTGCC-3

′

(

X = C or A, underline denotes the

mismatch). Samples were irradiated and electrophoresed through a 20% denaturing PAGE gel.

A light control (LC, without rhodium) and dark control (DC, without irradiation) were included.

A representative autoradiogram of a photocleavage titration with

rac

-Rh(bpy)

chrysi

(A,

arrows indicate positions of mismatched bases) and a representative sigmoidal curve fit of

pooled data from photocleavage titrations for binding constant determination (B) are shown.

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Figure 3.

Inhibitory effects of Rh(NH

)

chrysi

as a function of incubation time on cellular

proliferation. Shown are plots of BrdU incorporation (a measure of DNA synthesis and

therefore cellular proliferation) normalized to the BrdU incorporation of untreated cells as a

function of rhodium concentration. Standard error bars for five trials are shown. MMR-

proficient HCT116N cells (green) and MMR-deficient HCT116O cells (red) were plated and

allowed 24 h to adhere before incubation with 0–25

M Rh(NH

)

chrysi

for 12, 24, 48, or

72 h. At the end of the 12, 24, and 48 h incubations, the medium containing Rh was replaced

with fresh medium for the remainder of the 72 h, followed by ELISA analysis. BrdU was added

to the medium 24 h prior to analysis.

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Figure 4.

Inhibitory effects of rhodium metalloinsertors as a function of incubation time. Shown are plots

of BrdU incorporation normalized to the BrdU incorporation of untreated cells as a function

of rhodium concentration. The inhibition differential is the difference of the normalized

percentages of cellular proliferation for each cell line, with standard error bars (

N–O

√

(

)). ELISA analyses were performed as in Figure 3. Cells were incubated with no

rhodium, 2

rac

-Rh(DIP)2chrysi

, 10

M Rh(NH

)

chrysi

, or 20

rac

-Rh

(L)

chrysi

(L = HDPA, bpy, or phen).

Ernst et al.

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Figure 5.

Inhibitory effects of

rac

-Rh(DIP)

chrysi

. Shown are plots of BrdU incorporation normalized

to the BrdU incorporation of untreated cells as a function of rhodium concentration. Standard

error bars for five trials are shown. MMR-proficient HCT116N cells (green) and MMR-

deficient HCT116O cells (red) were plated and allowed 24 h to adhere before incubation with

0–5

rac

-Rh(DIP)

chrysi

for 12 h. At the end of the incubation, the medium containing

Rh was replaced with fresh medium, and cells were grown for an additional 60 h before ELISA

analysis. BrdU was added to the medium 24 h prior to analysis.

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Figure 6.

Inhibitory effects of rhodium metalloinsertors as a function of metal complex identity. Shown

are bar graphs of BrdU incorporation normalized to the BrdU incorporation of untreated cells

as a function of rhodium concentration. The inhibition differential is the difference of the

normalized percentages of cellular proliferation for the two cell lines, HCT116O versus

HCT116N. ELISA analyses were performed as in Figure 3. Cells were incubated with no

rhodium, 2

rac

-Rh(DIP)

chrysi

, 10

M Rh(NH

)

chrysi

, or 20

rac

-Rh

(L)

chrysi

(L = HDPA, bpy, or phen). A correlation between mismatch binding affinity and

differential inhibition of MMR-deficient cells is evident.

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Figure 7.

Crystal and model structures of rhodium metalloinsertors bound to the mismatch site. Rhodium

insertors (red) are shown bound to the DNA (gray) from the minor groove at the mismatch site

with the bases (adenine in blue, cytosine in yellow) ejected and the chrysi ligand stacked fully

with the adjacent base pairs. The crystal structure of

-Rh(bpy)

chrysi

bound to the CA

mismatch is shown in panel (A), along with structural models of

-Rh(DIP)

chrysi

(B) and

-Rh(NH

)

chrysi

complex upon the rhodium center of the bpy complex leads to steric clashes with the sugar–

phosphate backbone (possible atoms involved in green), whereas the tetraammine complex is

easily accommodated.

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Ernst et al.

Page 18

Table 1

Binding Affinities

of Rh(L)

chrysi

Complexes

for CC and AC Mismatches

Complex

Ancillary ligand

(CC)/M

-1

(AC)/M

-1

Rh(NH

)

chrysi

1.0 × 10

3.4 × 10

Rh(bpy)

chrysi

3.4 × 10

2.2 × 10

Rh(HDPA)

chrysi

2.0 × 10

2.6 × 10

Rh(phen)

chrysi

3.2 × 10

1.4 × 10

J Am Chem Soc

. Author manuscript; available in PMC 2010 February 18.