Towards a microparticle-based system for pooled assays of small

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Towards a microparticle-based system for pooled assays of small molecules in cellular contexts Carrie E. Yozwiak, Tal Hirschhorn, and Brent R. Stockwell ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/ acschembio.8b00043 • Publication Date (Web): 24 Jan 2018 Downloaded from http://pubs.acs.org on January 26, 2018

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ACS Chemical Biology

Figure 1: Schematic of bead-based, pooled screening technology

1 CELL 2 ID TAG 1 2 3 ID TAG S S S S 4 5 DEVD DEVD F 6 F Q Q 7 8 9 10 11 12 13 14 ID TAG 15 ID TAG 6 5 SH SH 16 17 Q 18 19 20 21 22 23 = Bead 24 = Fluorescent sensor 25 26 = Lethal small molecule 27 28 Q = Quencher of Fluorescent Sensor 29 30 DEVD = Caspase cleavable peptide sequence 31 S-S = Disulfide bond 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 ACS Paragon Plus Environment 60

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Figure 2 a

CellTiterGlo, Beads + HT-1080 cells 72 hour incubation

Page 2 of 34 Red fluorescence measurement, 72 h bead incubation in different solvents

1.0 MilliQ Water DMSO TFA Mes Buffer pH 6.3 Ethanol

Beads Beads-FITC Beads-C3Sensor

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2 4 6 10 10 10 535 nm Emission Reporter Fluorescence HT-1080 cells HT-1080 cells + beads

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Figure 3

1 b Design of Bead-FITC 2 a Design of Fluorescent Reporter, Bead-C3Sensor HO O 3 O O O O O 4 H H H H O N N N O O N N N NH O O H H 5 O O O O O O OH 6 NH S 7 O OH O HN 8 9 O 10 OH 11 25.54A 12 13 14 d Bead-C3Sensor Shows low 15 Reporter Fluorescence intensity 16 c Incubation of FRET peptides with active 17 caspase 3 protein 18 19 20 7 10 21 30 6 DEVD peptide 10 22 DEVG peptide 5 23 10 20 24 4 10 25 3 26 10 10 27 2 10 28 Bead-FITC 1 Bead-C3Sensor 29 10 0 Bead 0 30 5 30 60 120 180 10 31 0 2 4 6 Time (min) 10 10 10 10 32 530 nm Emission 33 Reporter Fluorescence 34 35 FRET gate established to sort 36 e f Caspase 3 protein incubation with Bead-C3Sensor Bead-C3Sensor from Bead-FITC population 37 shows increase in Reporter Fluorescnece 7 10 38 6 93+/- 2% population 39 10 FRET gate 40 5 10 41 4 7 42 10 10 19.7% cleaved population 43 6 3 10 FRET gate 10 44 5 2 10 45 10 46 4 1 Bead-C3Sensor 10 10 Bead-C3Sensor inside gate 47 0 3 10 10 48 0 2 4 6 2 49 10 10 10 10 10 50 Bead-C3Sensor 1 530 nm Emission 10 51 Bead-C3Sensor+Caspase 3 Reporter Fluorescence 0 10 52 0 2 4 6 53 10 10 10 10 54 530 nm Emission Reporter Fluorescence 55 56 57 58 59 ACS Paragon Plus Environment 60 Me N Me

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ACS Chemical Biology Figure 4

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

a H 2N

Design of barcode linker to bead (CH 2)12

O

O N H

NO 2 O O P OH O Oligonucleotide Barcode 3'

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PCR of barcodes linked to bead

FW Primer RV Primer Barcode 1 Barcode 2 Bead Bead Wash Supernatant 365 nm light irradiation

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PCR of Bead-Barcode after incubation with apoptotic cell lysate FW Primer RV Primer Barcode 2 365 nm light irradiation Live cell lysate Apoptotic cell lysiate

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Figure 5 a

Confocal image of Bead-FITC-Barcode incubated 6 h with HT-1080 cells i

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Z-stack slices show Bead-FITC-Barcode is internalized

Cell TiterGlo, 72 h HT-1080 incubation with system Normalized Viability

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ACS Chemical Biology Figure 6 b Bead- Malemide linker - DM1

a Bead- Disulfide linker - DM1 O

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Towards a microparticle-based system for pooled assays of small molecules in cellular

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contexts

3 Carrie E. Yozwiak1, Tal Hirschhorn2 and Brent R. Stockwell1, 2,*

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1

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2

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550 West 120th Street, New York, NY 10027, USA

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*Correspondence: [email protected]

Department of Chemistry, Columbia University, New York, NY 10027, USA Department of Biological Sciences, Columbia University, Northwest Corner Building, MC 4846,

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Keywords

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Small molecules; beads; pooled screening; microparticle; nanoparticle; screening; drug

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discovery

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ACS Chemical Biology 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Abstract

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Experimental approaches to the discovery of small molecule probes and drug candidates often

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use biochemical or cell-based screening of large libraries (>105) of small molecules. Small

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molecules of interest are tested one at a time in individual wells of a microtiter plate, at a

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significant cost in time and resources. Furthermore, evaluation of large numbers of compounds

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in such assays requires robust cellular or biochemical screening formats that may not be

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relevant to the contexts found in human patients. We envision a solution to these issues that

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involves a pooled system of small molecule screening, which would require development of

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numerous new technologies, and solutions to several key challenges. We report here that a

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microparticle-based screening system can allow for screening of small molecules in such a

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pooled fashion, analogous to the pooled screens of genetic reagents that have been powerfully

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deployed in recent years. We developed a cleavable linker that can link small molecules of

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interest to silica microparticle beads; a DNA tag encoding the identity of the small molecule on

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each bead that was attached to the silica beads through a photocleavable linker to enable its

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amplification; and a bead-based fluorescent sensor that can report on the activity of small

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molecules in cells. We suggest that this pooled small molecule screening system could

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ultimately be useful for drug and probe discovery, allowing rapid and inexpensive screening of

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small molecules in assays of relevance to human diseases.

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Introduction

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Therapeutic small molecule candidates often fail upon exposure to the complex physiology

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found in animal models and human patients, which are not captured in simple high-throughput

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tissue culture screening systems. In addition, the time and expense needed to assess the

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effects of large numbers of small molecules limits the number of compounds that can be tested,

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and requires significant resources.

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This problem of evaluating large numbers of reagents in physiologically relevant cell and animal

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models has been addressed for genetic reagents such as RNAi1, CRISPR2 and cDNA3, by

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creating barcoded retroviral libraries that can be used to infect target cells in culture or in animal

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models. Using these tools, effective reagents can be selected and decoded using a rapid and

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inexpensive procedure compared to testing of individual reagents one at a time in an arrayed

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fashion. In order to more efficiently analyze biological activities of small molecules, a pooled

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approach would similarly be useful. Large scale pooling of small molecules is not possible,

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because all cells are exposed to each compound, creating uninterpretable effects of large

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mixtures of compounds. This problem was solved for genetic reagents by physically packaging

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each test reagent into separate viral particles, which can deliver one genetic reagent to each

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infected cell, when the multiplicity of infection (MOI) is suitably calibrated.

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Pooling has thus emerged as a powerful approach to efficiently perform genetic screens. For

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example, pooled RNAi barcode screens provide a convenient method to evaluate large shRNA

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libraries. In this method, shRNA vectors are tagged with a unique oligonucleotide sequence

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before infection of cells, and then analyzed via PCR amplification and barcode sequencing. The

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understanding of gene function acquired by large-scale pooled RNAi screens has greatly

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contributed to understanding of cell signaling and drug target discovery for cancer and infectious

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diseases4-6. In addition, such pooled barcode shRNA screens have been performed both in cell



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culture assays and in animal models7,8.

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There is no pooled small molecule screening method for cellular assays that is analogous to

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pooled, genetic screening technology. A major limiting factor is that multiple small molecules

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can penetrate cells. The current paradigm is therefore a one-compound-in-one-well approach,

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which provides a straightforward analysis, at the cost of being relatively inefficient and

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expensive to screen large small molecule libraries without substantial automation9. Moreover,

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evaluating the effects of individual compounds in animal models requires great time, expense,

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and large numbers of animals.

70 71

Merrifield’s revolutionary development of solid-phase peptide synthesis set the stage for

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technologies that generate large libraries of diverse peptides and small molecules10. Split-pool

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synthesis methods rely on chemical conjugation of peptides or small molecules to a resin in a

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one-bead-one-compound (OBOC) format. After the initial coupling reaction is performed on

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different aliquots of beads, the beads are pooled and re-split, and a new chemical moiety is

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added to each group of beads. This cycle of split-pool synthesis is repeated to efficiently

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generate large libraries of structurally diverse compounds for screening11-13. Screening of these

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compound-bead libraries has been demonstrated in isolated protein systems, but not in intact

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cells, because cells generally do not internalize the synthesis beads used for split-pool

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synthesis14,15.

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Herein, we report several advances that enable the development of a technology for pooled

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small molecule screening in cellular assays. Using fluorescent silica microparticle beads coated

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with carboxylate groups, we covalently attached three components to the bead surface—(1)

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small molecules to be assayed via a cleavable linker, (2) a fluorescent reporter and a

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photocleavable barcode encoding the identify of each test small molecule. We examined the


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utility of each aspect of this system in proof of concept experiments in human tumor cell culture

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models.

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Results

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We designed a microparticle-bead-based system that could deliver small molecules into cells

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and report on the identity of the small molecule tested and its cellular activity (Figure 1). Once

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taken up into cells, the beads were designed to release each test small molecule either by

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disulfide bond cleavage or cleavage of a linker peptide by cellular proteases. We envisioned

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that a fluorescent reporter would indicate if the small molecule delivered to a particular cell was

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exerting the biological activity of interest. For example, if a small molecule initiates apoptosis,

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executioner caspase 3 would be activated; caspase 3 cleaves the DEVD amino acid

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sequence16; other fluorescent peptide reporters could similarly report on the activity of other

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proteases in cells. A fluorescent reporter attached to beads was thus designed to detect

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caspase 3 activation; it consists of a fluorescence resonance energy transfer pair 17 separated

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by a DEVD peptide sequence. Once apoptosis occurs as a result of exposure of the cells to the

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released small molecule, activated caspase 3 cleaves the DEVD sequence, resulting in a loss of

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FRET. The increase in the fluorescence intensity of the donor fluorophore can be detected by

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flow cytometry, and the bead can then be separated from beads in which the FRET signal

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persists. Finally, a cleavable oligonucleotide barcode attached to the beads can be amplified

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using PCR and then sequenced, to identify the small molecule that caused caspase activation in

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the cellular context being tested (Supplementary Figure 1).

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We reasoned that during the course of a pooled small molecule screen, the beads must be

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harvested in order to sequence barcodes that encode the identity of the test compounds

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attached to each bead. The beads must thus be made of a material that does not affect cell

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viability, and that does not decompose upon cellular internalization or cell lysis. Cell viability



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experiments with varying concentrations of 1 µm silica microparticles showed no lethality after

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incubation for 72 h (Figure 2a), suggesting these beads are suitable, based on these criteria.

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Microparticle fluorescence and structure were maintained after incubation in dimethylsulfoxide

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(DMSO), dimethylformamide (DMF), ethanol, water, and 2-(N- morpholino)ethanesulfonic acid

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(MES) solvents, suggesting that silica beads are compatible with a variety of organic synthesis

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conditions, in contrast to other solid-phase microparticle materials we examined. Regardless of

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the solvent environment, the fluorescence intensity of the beads at the appropriate emission

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detection channel of a flow cytometer remained constant (Figure 2b).

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Dynamic light scattering (DLS) was used to analyze microparticle size distribution profile of

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beads (Figure 2c). DLS measurements assess the scattering of a monochromatic light source

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by particles in a solution; the similar size distribution of each group of beads demonstrated bead

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stability across diverse solvents. Additionally, we found that the carboxylate functionality on

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these beads was compatible with bioconjugation reactions with oligonucleotides and peptides.

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We thus envisioned that silica particles would act as the delivery vehicle for the system. Given

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the robust solvent stability of silica microparticles and their compatibility with diverse synthetic

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organic chemistry reaction conditions, we next examined the biocompability and size

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requirements of this material. We realized that the beads must be small enough to be

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internalized by cells, yet must possess a sufficient concentration of synthetic handles to

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conjugate a suitable amount of each component onto the bead. Our goal was to find a bead

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type that demonstrates sufficient cellular uptake with a maximum of functional handles for

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chemical synthesis. To evaluate the appropriate balance of these factors, we studied cellular

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uptake of fluorescent microparticles that were 1 µm in diameter in HT-1080 fibroscaroma cells.

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Because the beads possess a fluorescence emission maximum at 625 nm, cells that


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internalized the beads showed an increase in fluorescence on the FL2 channel of the flow

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cytometer, which detects a fluorescence emission of 585/40nm (Figure 2d). The fluorescence of

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10,000 live HT-1080 cells without beads (n = 4) was examined as a control to develop a suitable

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gate on the bead emission detection channel of the flow cytometer versus the fluorescence

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emission of the reporter component (FITC, excitation 488 nm, emission recorded on 585 nm

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channel) such that the gate encompassed 99.8 +/- 0.2% of the population. The gate was

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applied to each sample, and was used to discriminate between cells without beads versus cells

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with fluorescent beads. Of the live cell population analyzed, 8.2 +/- 0.4% of cells did not take up

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a 1 µm bead after six-hour incubation (n = 3); this population did not show an increase in

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fluorescence intensity on the 585 nm channel, the bead emission fluorescence channel. We

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found that smaller beads were taken up by virtually all cells, but the surface area available and

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fluorescence intensity on smaller beads were less favorable for compound delivery

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(Supplementary Figure 2a). Confocal microscopy confirmed cellular internalization of these 1

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µm diameter beads (Figure 2e); thus, we determined that this size would be ideal for maximizing

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available microparticle surface, while still being amenable to cell uptake. By adapting the

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multiplicity of infection concept from viral screening, we were able to deliver a single bead into

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individual cells (Supplementary Figure 2b). 1 µm silica beads were internalized by multiple

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different human cancer derived cell lines (Supplementary Figure 2c), suggesting these beads

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would be suitable for examining biological activity in a variety of contexts.

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A FRET (Fluorescence Resonance Energy Transfer) sensor can act as a reporter of protease

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activity, to allow for selection of compounds that induce specific protease activity18. The integrity

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of the FRET pair should be maintained in live cells and disrupted upon activation of a protease

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to yield a detectable change in fluorescence. We evaluated this concept initially with small

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molecules that cause cell death through caspase-dependent apoptosis, a well-defined

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mechanism of cell death with biomarkers available for reliable detection19. Additionally, the



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evasion of apoptosis is a hallmark of cancer, so discovering small molecules that initiate

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apoptosis might be therapeutically relevant; indeed, venetoclax, an inhibitor of Bcl-2 proteins

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that induces apoptosis, has recently been approved for clinical use by the U.S. FDA20.

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Caspase 3 is the main executioner enzyme in apoptosis21,22. This cysteine protease specifically

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cleaves after the peptide sequence DEVD16,22. For effective fluorescence quenching, FRET

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relies on three principles: (1) overlap between the emission spectrum of the donor fluorophore

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and the excitation spectra of the acceptor fluorophore, (2) a donor-acceptor pair distance of 1-

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10 nm, and 23 suitable orientation of the FRET pair for effective non-radiative, resonance

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transfer of energy24. Because FRET is a distance-dependent effect, we placed the DEVD

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substrate between the FRET pair to create ‘C3sensor’ (Figure 3a). Upon caspase 3 activation,

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the peptide should be cleaved, to increase the distance from the donor fluorophore (FITC) to the

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acceptor chromophore (dabcyl). FRET pair separation should result in increased fluorescence

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signal of the FITC dye (Figure 3b). Indeed, we found that the fluorescence intensity of FITC

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increased 15-fold when incubated with 0.5 units of purified active caspase 3 protein for 2 hours

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(Figure 3c).

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The FRET-caspase sensor was conjugated to silica microparticles by an amide linkage via 1-

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ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) coupling of the primary amino terminus to

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the carboxylate-bead coating. We designated this covalently linked reporter-bead complex as

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‘Bead-C3Sensor.’ To ensure that the peptide was covalently bound to the bead, we analyzed

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the zeta potential of the microparticles. Zeta potential measurements assess the electrostatic

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potential of the surface of particles suspended in aqueous solution by tracking the movement of

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charged particles across an applied electric field25. Similarly, charged particles in solution

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correlate to a large zeta potential, demonstrate stable colloidal dispersion, and low

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agglomeration of the suspension. After conjugation, the zeta potential increased, demonstrating


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less repulsion between particles and thus supporting effective conjugation (Supplementary

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Figure 3a). To examine cellular internalization of the microparticle-peptide system, a peptide

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labeled with fluorescent FITC without the dabcyl quencher was used as a control (Figure 3b).

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The FITC-only containing peptide is equivalent to the cleaved product of the bead with the

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FRET pair exposed to caspase 3, and shows high fluorescent intensity at two emission peaks

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corresponding to both the bead and the dye (Figure 3d). This peptide fluorescence would be

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equivalent to 100% protease efficiency in cleaving each conjugated DEVD sequence.

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Fluorescence measurements demonstrated covalent attachment of the FITC-peptide to the

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bead (Supplementary Figure 3b), and confocal microscopy showed internalization of the

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peptide-bead system, Bead-FITC (Figure 5a).

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We recognized that it is essential to distinguish the fluorescent reporter in the quenched FRET

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pair on beads from beads exposed to active caspase 3, so that flow cytometry can effectively

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separate each population. Ineffective separation would result in a high degree of false-positives.

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Flow cytometry analysis of the Bead-C3Sensor conjugated pair showed comparable FITC

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fluorescence intensity to the bead alone (Figure 3d). Both of these populations were distinct

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from the bead-FITC conjugated pair, which showed a distinct increase in fluorescence intensity

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on the reporter fluorescence detection channel (Figure 3d). After the Bead-C3Sensor was

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exposed to active purified, active caspase 3 protein, there was a distinct shift to higher reporter

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emission fluorescence intensity when compared to the original, uncleaved FRET state (Figure

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3f). When a gate that encompasses 93 +/- 2% of the native Bead-C3Sensor system (Figure 3e)

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was applied to the Bead-C3Sensor exposed to active caspase 3 protein, 70 +/- 7 % of the

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beads have increased reporter fluorescence sufficient to fall outside of the gate (Figure 3f). That

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population is the target for harvesting, and the barcode of those samples could be sequenced to

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identify apoptosis-inducing small molecules.

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Furthermore, we tested whether a fluorescent reporter could be designed to specifically sense

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the activity of other proteases in a cellular context. We thus replaced the caspase-sensitive,

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DEVD peptide motif in the Bead-C3Sensor with a cathepsin D cleavable motif (bead-

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CatSensor). Cathepsin D is a lysosomal aspartic endopeptidase localized in intracellular

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structures, such as endosomes, lysosomes, or phagosomes; as such, it should co-localize with

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the bead system if the beads are internalized by endocytosis.

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We reasoned that cathepsin D access to the bead-sensor system would cleave the peptide

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motif, separate the FRET pair, and increase sensor fluorescence. Indeed, we found that HT-

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1080 cells containing the bead-CatSensor exhibited a 3.6-fold increase in sensor fluorescence

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over cells containing the bead control (Supplementary figure 3c). Additionally, the Bead-

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CatSensor system exhibited a 70% increase in fluorescence intensity in HT-1080 cells

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compared to cells treated with chloroquine, which should suppress lysosomal cathepsin D

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activity. Finally, we detected a 1.5-fold increase in fluorescence intensity in cells with this sensor

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compared to cells pre-treated with the cathepsin inhibitor Pepstatin A, suggesting that we were

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reporting on cathepsin protease activity. Together, these experiments suggest that it is possible

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to design protease sensors that report on different protease activities in cells, particularly in the

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lysosomal compartment where the beads get taken up.

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Next, we recognized that each barcode must be uniquely matched to each small molecule

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tested, show no bias in PCR amplification, stay covalently bound to the bead throughout the

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assay, and be sequenced to reveal a statistical distribution of ‘hits’. We developed

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oligonucleotide barcodes consisting of 45 unique base pair sequence flanked by forward and

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reverse primer regions (Supplementary Table S2). Each distinct barcode sequence used the

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same set of primers, so multiple ‘hit’ small molecules out of the pool could be equally amplified


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and identified. The primer design includes an overhang region with a sequence compatible with

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existing DNA sequencing technology.

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Prototype DNA barcodes were synthesized and attached to beads through an amide linkage

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after reaction with a terminal primary amine on the 5’ end. In addition, a photocleavable linker

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was included in the barcode design (Figure 4a), which we found necessary for acceptable PCR

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efficiency. Upon irradiation with near-UV light (300 – 350 nm range to avoid DNA damage),

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cleavage of the DNA from the bead was observed after 10 minutes (Fig 4b). This cleavable

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linker was included to make PCR amplification of the oligonucleotide more efficient after

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isolation of the bead, as we found that PCR amplification was inefficient on beads: lack of

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amplification was observed when the oligonucleotide remained bound to the bead and can likely

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be explained by steric hindrance of the bead to the polymerase. The bulky bead may block the

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polymerase from binding to the oligonucleotide, preventing DNA extension.

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Two unique barcodes were initially used for a proof of principle to ensure effective primer

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binding to more than one sequence (Figure 4b). Once the barcode was conjugated to the bead,

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the system was incubated for 24 h with the lysate of cells that were undergoing apoptosis.

259

Despite protease activation and other environmental changes that occur during the programmed

260

death pathway, the oligonucleotide was effectively PCR amplified after photo cleavage from the

261

bead (Figure 4c). This suggests the bead-barcode conjugated pair can persist throughout the

262

entirety of the screen for small molecule identification, a key feature of a pooled small molecule

263

screening system.

264 265

To evaluate the compatibility of the sensor and the barcode, both were covalently coupled to the

266

bead by a one-pot EDC coupling, as confirmed by zeta potential measurements. The addition of

267

the oligonucleotide to the bead-peptide conjugate made the zeta potential slightly more negative



268

(Supplementary Figure 3a)). Additionally, dynamic light scattering showed an increase in

269

particle size upon barcode attachment (Supplementary Figure 4). Confocal microscopy showed

270

internalization of this bead-dye-oligo reagent (Figure 5a and b), and that this was not lethal to

271

HT-1080 cells after 48 h incubation (Figure 5c).

272 273

An additional key component of the small molecule pooled screening technology needed is

274

covalent attachment of test small molecules. The small molecules of interest should detach from

275

the bead only once the system is internalized into cells. We first examined the utility of the

276

potentially glutathione-reductase-rich environment of the cytosol to employ a disulfide-bond

277

linker to connect the bead to the small molecule (Supplementary Figure 5). Although a covalent

278

bond is maintained outside of the cellular environment, once internalized, the disulfide bond

279

should be reduced to cleave the small molecule from the cell. Once inside the cell, the small

280

molecule should not contain any additional chemical modifications that can alter its biological

281

activity. The lethal small molecule used for this proof of concept experiment was the microtubule

282

inhibitor mertansine (DM1). DM1 was chosen because of its potency (4.6 nM in HT-1080 cells)

283

to induce apoptosis (Supplementary Figure 6b). Additionally, DM1 possesses a terminal thiol

284

moiety in its active state, thus simplifying the model by eliminating the need for a self-immolative

285

linker. The chemistry used to covalently attach the known lethal molecule is outlined in

286

Supplementary Scheme 1.

287 288

The first step of integrating the components of the system involves differentiating a portion of

289

the surface of the bead with a functionality to react with the small molecule. We sought a high

290

concentration of each small molecule to be bound to each bead, to maximize the ultimate

291

cellular concentration achieved. When using 0.5 equivalents of mono-Boc protected diamine per

292

one equivalent of carboxylate group on the bead surface, there is a large enough concentration

293

of unreacted carboxylate handles on the bead for effective conjugation and detection of the


Page 18 of 34

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294

barcode and fluorescent reporter, but the majority of chemical space on the bead surface will be

295

unreactive to these moieties, having been coupled to the protected diamine. It is thus inert to the

296

subsequent EDC couplings and reserved for small molecule attachment. Deprotection of the

297

Boc moiety to reveal a primary amine was possible with concentrated trifluoroacetic acid

298

because the silica bead withstands strong acid and the more sensitive groups (peptide,

299

oligonucleotide) not yet attached (Figure 2c). Facile coupling of 3-(2-pyridyldithio)propionic acid

300

N-hydroxysuccinimide ester (SPDP) to the newly differentiated bead surface yields a pyridine-

301

disulfide moiety that withstands the chemistry to attach the oligo and fluorophore, yet will react

302

during a simple and robust thiol-sulfide exchange reaction to attach the lethal small molecule to

303

the bead in the last step (Figure 6a). Based on the average cell diameter (20 microns), the

304

reaction efficiency, and the number of carboxylate groups per bead, we have calculated the

305

average cellular concentration of the small molecule to be ~100 µM, which is more than

306

sufficient to identify promising compounds out of a first-pass screen, even if overall loading and

307

release efficiency is low. Based on the coupling equivalents, this could be tuned to lower

308

concentrations for more stringent assays.

309 310

To test the actual release of an active small molecules in cells, we synthesized a non-cleavable

311

linker using malemide bioconjugation chemistry to control for disulfide bond reduction and

312

release of the small molecule after bead uptake (Figure 6b). We did not detect apoptosis with

313

either cleavable or non-cleavable small molecule-bead conjugated system, suggesting cellular

314

reductases could not release the compound from the disulfide linker; this suggests that the

315

beads do not get released from the lysosome to the cytosol.

316 317

We thus evaluated the cathepsin D cleavable linker, which should release compounds from

318

endosomal compartments when the bead system encounters cathepsin D. We conjugated the

319

lethal molecule monomethyl auristatin E (MMAE) to beads via a cathepsin D cleavable linker



320

(Figure 6c). After incubation of this bead system with HT-1080 cells, we observed cell lethality

321

only when the cleavable compound was present (Supplementary Figure 6a), suggesting that

322

cathepsin-cleavable linkers are more effective for small molecule release, compared to disulfide

323

linkers (Figure 6d) in the current system. Together, these constitute key elements needed for a

324

pooled small molecule screening system, involving a solid phase system that can be taken up

325

by cells, a cleavable linker that gets released upon cellular uptake, a fluorescent reporter that

326

can indicate compound activity, and an amplifiable barcode that can specify compound identity.

327 328

Discussion

329

Pooled screening of small molecules offers efficient biological evaluation of large compound

330

libraries, and thus has distinct advantages over traditional high throughput screening methods.

331

The potential to select small molecules that modulate biological activity through first-pass

332

screens in vivo would expand current drug discovery capabilities. We examined each individual

333

component needed for pooled, small molecule screening, whereby active small molecules could

334

be conveniently identified based on fluorescent reporter activity.

335 336

We identified a suitable solid phase upon which to develop a small molecule delivery system for

337

pooled screening: fluorescent silica microparticles provided enough sites for chemical

338

diversification and maintained their inherent qualities throughout organic chemical reaction

339

conditions. Additionally, their fluorescence allows for easy particle tracking throughout the assay

340

and convenient distinction from cellular debris for harvesting upon assay completion.

341 342

We designed fluorescent sensors of small molecule activity. When conjugated to beads, such

343

sensors show a distinct signal after exposure to enzymatic activity specific to apoptotic cell

344

death. Additionally, such sensors can likely be tuned to detect enzymatic activity of diverse

345

peptidases.


Page 20 of 34

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346 347

Additionally, we established a barcode system to report on the identity of each small molecule.

348

The barcode is compatible with deep sequencing technology, so that a statistical distribution of

349

small molecules can be determined from the pooled assay, thus reducing the ‘false positive’ hit

350

molecules from the screen. The barcode does not affect the ability of the bead to be taken up by

351

HT-1080 cells, stays with the bead throughout the duration of the assay, and can be amplified

352

after incubation with cells undergoing death.

353 354

We initially designed this system discover lethal small molecules that specifically initiate cell

355

death by caspase-3. We envision that the system can be tuned and applied to wide-ranging,

356

diverse cell models and biochemical processes as a drug and probe discovery tool. The peptide

357

linker of the reporter can be modified to report on other proteases, protein-protein interactions,

358

transcription, or other biological events. We tested the feasibility of doing this by changing the

359

sensor to detect cathepsin D. Because the data suggest that this bead system is trafficked into

360

the cell via endocytic pathways, the beads appear to be subject to endosomal entrapment. In

361

addition to investigating strategies to facilitate escape from the endosome, one can use

362

endosomal localization to detect activities in lysosomes and endosomes.

363 364

Endosomal/lysosomal targeting is one application of this technology, as there are numerous

365

diseases associated with the disruption of lysosomal function, including Tay-Sachs disease,

366

Gaucher’s disease, Hurler syndrome, among others. Often, low enzymatic activity in the

367

endosome or lysosome contributes to these disorders; this screening system enables the cell-

368

based screening for small molecules that enhance lysosomal protease activity.

369 370

The conjugation of mertansine to the bead system requires thiol-sulfide exchange chemistry, but

371

covalent attachment of the small molecule to the bead was effective using maleamide chemistry



372

or NHS-reactive linkers to expand the scope of reactive moieties on the small molecule. In

373

addition, functional handles can be broadened to include click techniques, alcohols, etc. The

374

system can be synthesized such that a user can acquire a specific enzyme

375

sensor/barcode/bead system, and then simply conjugate to their small molecules of interest.

376

The ability to use simple, robust bioconjugation chemistry to conjugate diverse linkers to the

377

bead system enhances the kinds and numbers of small molecules that can be screened in this

378

type of system.

379 380

To address applicability to a broader chemical library, we explored the usage of a self-

381

immolative linker to attach the small molecule via the disulfide bond. After the disulfide bond is

382

reduced, the terminal thiol can participate in a nucleophilic substitution reaction with the

383

carbamate to form a five-membered ring; this releases the amine-containing small molecule, in

384

its original state, without any modification. Small molecules included in the library include a

385

reactive handle by which they are conjugated, which is a current limitation.

386 387

These studies were designed to provide an initial proof of concept. While this disulfide system

388

was not effective at releasing compounds under the conditions we examined, it might be

389

possible to engineer cells to overexpress specific reductases that can cleave such a linker. We

390

also found that a cathepsin-cleavable linker was more readily cleaved in HT-1080 cells,

391

suggesting that this may be a simpler release mechanism. Ultimately, further optimization and

392

diversification of the components of the system may have numerous applications drug discovery

393

and allow for diverse pooled small molecule screens.

394 395

Methods

396

Bead incubation with cells.


Page 22 of 34

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397

Red (540/625 nm) fluorescent, non-magnetic, monodisperse, carboxylate coated screenCORE

398

microspheres were purchased from Chemicell (Berlin, Product Number 6101-1) HT-1080

399

(fibrosarcoma) cells were obtained from American Type Culture Collection (Manassas, VA,

400

ATCC CCL-121). HT-1080 cells were grown in DMEM High-glucose medium supplemented with

401

1% non-essential amino acids (Mediatech, Product Number 10-013-CM) 1% P/S, and 10%

402

heat-inactivated fetal bovine serum (FBS). All the cell lines were grown at 37 °C under 5% CO2

403

and harvested at 80% confluence for experiments. Uptake experiments were completed in 6

404

well plates (Corning costar (Corning, NY, 3516). 200,000 HT-1080 cells were plated per well

405

and let adhere overnight at 37 °C under 5% CO2. After six-hour incubation with beads at 37 °C,

406

the cells were washed twice with PBS, harvested with 0.25% Trypsin-EDTA (1X) Phenol red

407

(Invitrogen (Carlsbad, CA, 25200-114)), re-suspended in growth medium, and strained by a 70

408

µm filter (Fisher Scientific, 352350) into Eppendorf tubes for flow analysis.

409 410

Technology synthesis

411

All chemicals were obtained from Sigma Aldrich and used without further purification. All

412

peptides were synthesized by LifeTein (Somerset, NJ). Oligonucleotides were synthesized by

413

Integrated DNA Technologies (Coralville, IO). Peptide and oligonucleotide sequences are

414

detailed in Supplementary Tables 1 and 2. This procedure was performed open to the

415

atmosphere. 20 µL beads (Chemicell (Berlin, Product Number 6101-1)) were washed with 0.1

416

M MES buffer (pH 6.3) by centrifugation at 14800 rpm (16162 xg) for 5 min, removing the

417

supernatant, re-suspending in buffer, and centrifuging again. The bead pellet was re-suspended

418

in 85 µL N-Hydroxysulfosuccinimide sodium salt (sulfo-NHS) (20 mM in MES buffer), 8.5 µL N-

419

Boc- 2,2′-(ethylenedioxy)diethylamine (50 mM, 0.5 equiv to carboxylate on bead surface), then

420

42.5 µL N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide (EDC) (100 mM in MES buffer). The

421

reaction was stirred overnight at 37 °C before Boc-deprotection with 1000 µL 1:1 water:

422

trifluoroacetic acid. After 30 minutes, the reaction was washed six times by centrifugation and



423

bead re-suspension as described above. To the differentiated beads, 3-(2-pyridyldithio)propionic

424

acid N-hydroxysuccinimide ester (SPDP, 17 µL, 10 mM) was coupled by addition of 42.5 µL

425

EDC (100 mM in MES buffer). The reaction was allowed to stir overnight at 37 °C before

426

washing six times. The bead pellet was re-suspended in 85 µL N-hydroxysulfosuccinimide

427

sodium salt (sulfo-NHS) (10 µM in MES buffer), 0.68 µL peptide (5 mM, 0.001 equiv to

428

carboxylate on bead surface), 18 µL oligonucleotide (0.1 mM solution) then 42.5 µL N-(3-

429

dimethylaminopropyl)-N′-ethylcarbodiimide (EDC) (100 mM in MES buffer). The reaction was

430

shaken at 37 °C overnight before washing five times with water, then four times with 0.5%

431

Triton-X. Fluorescence of the supernatant of the final washing (494 nm excitation, 525 nm

432

emission) was recorded to ensure there no unconjugated fluorophore remained with the beads.

433

PCR reactions (procedure described below) of the final washing supernatant were completed to

434

ensure no uncoupled barcode remained. Excess mertansine (DM1, Medchem Express,

435

Monmouth Junction, New Jersey, Serial number G01432) (10 mM in DMSO, 17 µL) was added

436

to the bead pellet for final coupling, shaken at 37 °C overnight before washing five times with

437

DMSO. The final system was suspended in 20 µL DMSO (1 µL DMSO per 1 µL bead from

438

stock solution).

439 440 441

Synthesis of Bead-Malemide-DM1 control

442

All chemicals were obtained from Sigma Aldrich and used without further purification. This

443

procedure was performed open to the atmosphere. 20 µL beads (Chemicell (Berlin, Product

444

Number 6101-1)) were washed with 0.1 M MES buffer (pH 6.3) by centrifugation at 14800 rpm

445

(16162 xg) for 5 min, removing the supernatant, re-suspending in buffer, and centrifuging again.

446

The bead pellet was re-suspended in 85 µL N-Hydroxysulfosuccinimide sodium salt (sulfo-NHS)

447

(20 mM in MES buffer), 8.5 µL N-Boc- 2,2′-(ethylenedioxy)diethylamine (50 mM, 0.5 equiv to

448

carboxylate on bead surface), then 42.5 µL N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide


Page 24 of 34

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449

(EDC) (100 mM in MES buffer). The reaction was stirred overnight at 37 °C before Boc-

450

deprotection with 1000 µL 1:1 water: trifluoroacetic acid. After 30 minutes, the reaction was

451

washed six times by centrifugation and bead re-suspension as described above. To the

452

differentiated beads, N-ß-maleimidopropyl-oxysuccinimide ester (BPMS, Thermo Fisher

453

Scientific, Grand Island, NY, Catalog number 22298, 17 µL, 10 mM) was coupled by addition of

454

42.5 µL EDC (100 mM in MES buffer). The reaction was allowed to stir overnight at 37 °C

455

before washing six times. Excess mertansine (DM1, Medchem Express, Monmouth Junction,

456

New Jersey USA, Serial number G01432) (10 mM in DMSO, 17 µL) was added to the bead

457

pellet for final coupling, shaken at 37 °C overnight before washing five times with DMSO. The

458

final system was suspended in 20 µL DMSO (1 µL DMSO per 1 µL bead from stock solution).

459 460

Synthesis of Bead-Cathepsin D cleavable linker- monomethyl auristatin E

461

All chemicals were obtained from Sigma Aldrich and used without further purification. This

462

procedure was performed open to the atmosphere. 20 µL beads (Chemicell (Berlin, Product

463

Number 6101-1)) were washed with 0.1 M MES buffer (pH 6.3) by centrifugation at 14800 rpm

464

(16162 xg) for 5 min, removing the supernatant, re-suspending in buffer, and centrifuging again.

465

The bead pellet was re-suspended in 85 µL N-hydroxysulfosuccinimide sodium salt (sulfo-NHS)

466

(20 mM in MES buffer), 8.5 µL Boc- 2,2′-(ethylenedioxy)diethylamine (50 mM, 0.5 equiv to

467

carboxylate on bead surface), then 42.5 µL N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide

468

(EDC) (100 mM in MES buffer). The reaction was stirred overnight at 37 °C before Boc-

469

deprotection with 1000 µL 1:1 water: trifluoroacetic acid. After 30 minutes, the reaction was

470

washed six times by centrifugation and bead re-suspension as described above. The

471

differentiated beads and 42.5 µL EDC (100 mM in MES buffer) were added to a solution of 3-

472

Mercaptopropionic Acid (17 µL, 10 mM) and 85 µL N-Hydroxysulfosuccinimide sodium salt

473

(sulfo-NHS) (20 mM in MES buffer). The reaction was allowed to stir overnight at 37 °C before

474

washing six times. The beads were re-suspended in 50 µL DMSO and excess



475

maleimidocaproyl-L-valine-L-citrulline-p-aminobenzyl alcohol p-nitrophenyl carbonate (MC-Val-

476

Cit-BAP-PNP) (Accela Chemicals, San Diego, California, Compound ID SY034490) was added

477

to the reaction. The reaction was shaken at 37 °C overnight before washing five times with

478

DMSO. To the beads was added excess monomethyl auristatin E (MMAE, Medchem Express,

479

Monmouth Junction, New Jersey USA, Catalog number HY-15162) in DMSO. The reaction was

480

shaken at 37 °C overnight before washing five times with DMSO. Supernatant from the final

481

wash was kept for cell viability analysis.

482 483

Bead characterization

484

Microparticle size and zeta potential were measured on a Malvern Zetasizer Nano ZS (Malvern,

485

United Kingdom). For all measurements, microparticles were diluted 1:1000 in MilliQ water at

486

neutral pH. The reported diameters are the average of three measurements, where each

487

measurement comprises at least 10 acquisitions. The zeta potential was calculated according to

488

the Smoluchowski approximation26.

489 490

The absorbance of bead-FITC-coupling reaction supernatant was measured at 494 nm to

491

assess peptide coupling efficiency. Concentration was calculated using the Beer-Lambert law27.

492

% Coupling efficiency = ([FITC Reaction] – [FITC Supernatant]) / [FITC Reaction] x 100.

493 494

Biochemical assay for FRET detection.

495

0.5 units of purified, active caspase 3 protein (Abcam (Cambridge, MA, ab52101)) were

496

incubated with 2 µL of beads in 100 µL Apo-One buffer (Promega (Madison, WI, G7791)) for

497

two hours at 37 °C. The beads were washed twice with MilliQ water and re-suspended in MilliQ

498

water for flow cytometry analysis.

499 500

Flow Cytometry experiments.


Page 26 of 34

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501

Flow cytometric analysis was assessed on an Accuri C6 Flow Cytometer with a 488 nm Laser.

502

The 530/30nm filter (FL1) was used for FITC discrimination and the 585/40nm filter (FL2) was

503

used for bead identification. Using a live cell gate on DMSO treated cells, 10,000 events were

504

recorded for each population. FACS sorting was accomplished at the Herbert Irving

505

Comprehensive Cancer Center using a BDFacsAria II Cell Sorter.

506 507

Cell viability experiments

508

Assay plates were prepared by adding 1,000 HT-1080 cells per well in 35 µL of growth media to

509

opaque white 384 well plates (PerkinElmer (Waltham, MA, 6007688). After adhering to the plate

510

overnight, growth medium was removed from the cells and replaced with 35 µL lethal molecule

511

or beads in a two-fold dilution series. After 24 hours in the incubator, CellTiter-Glo Luminescent

512

Cell Viability Assay (Promega (Madison, WI, G7573) was completed. Percent viability was

513

calculated using cell luminescence intensity values determined from a Victor 3 plate reader

514

(PerkinElmer).

515 516

Confocal fluorescence imaging

517

Coverslips with HT-1080 cells incubated with beads (procedure described above) were washed

518

with PBS and fixed with 4% paraformaldehyde at room temperature for 10 minutes. After 3

519

washes with PBS, the cover slips were mounted on slides using ProLong Diamond Antifade

520

Mountant with DAPI (Thermo Fisher (Rochester, NY, P36966)). Confocal images were taken

521

with a Zeiss AxioOberever LSM700 at 63x and the Z Stacks were compressed with Zen

522

software. Analysis was done using ImageJ (National Institutes of Health, Bethesda MD). All

523

pictures are single confocal section imaged eight to twelve times to reduce noise.

524 525

Barcode detection



526

Beads were exposed to 365 nm light for 10 minutes for PC linker cleavage. The beads were

527

centrifuged at 14800 rpm for 5 minutes, and 1 µL of the supernatant was removed for PCR

528

amplification. Phusion Hi-Fidelity DNA polymerase (New England Biolabs) was used with

529

primer pair CEY_TruSeq_RPI1: CEY_TruSeq_RP1 (Sequence provided in Supplementary

530

Data).

531 532

Figure Legends

533

Figure 1. Design of bead-based system for pooled screening of small molecules. (1) The

534

technology, containing the covalently bound small molecule, identification tag, and activity

535

reporter, is internalized by the cell. (2) Once inside the cell, the small molecule is released

536

through the reduction of the disulfide bond linker. 23 The small molecule exists inside the cell

537

without chemical modification. (4) If the small molecule causes apoptosis, activated caspases

538

will cleave the DEVD peptide sequence bridging FRET pair. (5) Loss of FRET results in

539

increased fluorescence on the bead. (6) The activated system will be sorted based on this

540

change in fluorescence, and the barcode sequenced to identify the small molecule that caused

541

cell lethality. Red bead, silica microparticle; S-S, disulfide linker; blue square, lethal small

542

molecule; Q, quencher; F, fluorescent dye.

543 544

Figure 2. Selection of 1 µm silica bead for technology transport vehicle. (a) Cell viability of

545

red beads after 72 h incubation with HT-1080 cells shows no lethality. (b) Flow cytometry data

546

of beads show diverse solvents do not alter bead fluorescence detected on the 585/40nm

547

emission detection (FL2) channel. (c) Dynamic Light Scattering (DLS) analysis of beads after 72

548

h incubation with diverse solvents shows no bead degradation. (d) Scatter plot of flow cytometry

549

data. A gate of the live cell population before bead incubation was applied, and 10,000 events

550

for each trial were recorded inside the gate. 92 +/- 0.40% of HT-1080 cells incubated with 1 µm

551

diameter bead showed a 99-fold average increase in 585/40 nm filter (FL2 detectable


Page 28 of 34

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552

fluorescence) over control population after 6 hours. (e)Confocal Z-stack image of HT-1080 cells

553

incubated for 6 hours with 1 µm diameter silica beads show microparticle uptake. (i)Red

554

bandpass filter shows fluorescent bead. (ii) Green bandpass filter shows no bead bleadthrough.

555

(iii) Transmitted light. (iv) DAPI. (v) Overlay

556 557

Figure 3. Design and activity analysis of fluorescent reporter component. (a) Design of

558

FRET-based reporter conjugated to the bead, which we refer to as ‘Bead-C3Sensor’. Upon

559

caspase 3 exposure, the DEVD sequence is cleaved after the aspartic acid to release the

560

dabcyl quencher. FITC fluorescence increases, resulting from the distance-dependent loss of

561

FRET. (b) Design of the Bead-FITC system, used as a control for cell intake studies and flow

562

cytometry analysis. (c-e) Biochemical assay to assess FRET efficiency. (c) Flow cytometry data

563

of the bead without peptide (red), the bead conjugated to FRET peptide: Bead-C3Sensor (dark

564

blue) and the bead conjugated to the FITC peptide: Bead-FITC (light blue). 10,000 events were

565

recorded for 7 different samples of each population. The bead emission detection signal (y-

566

axis) stayed consistent between all populations, demonstrating the bead fluorescence was not

567

altered. The reporter emission detection channel (x-axis) registers the FITC signal, and was

568

quenched upon dabcyl addition. F test between FITC and FRET populations on the reporter

569

emission detection channel = 0.26 (n = 7). (d) The FRET gate for FACS sorting. A gate 28 was

570

included to encompass 93+/- 1.8% of the Bead-C3Sensor population (n = 8). (e) The FRET gate

571

28

572

average, the gate overlaps with 30 +/- 7.0% of the population (n = 10). The remainder of the

573

sample (blue) will be harvested for barcode sequencing. On average, we can expect to

574

sequence 70% of the lethal small molecule population. Overlay of the Bead-C3Sensor system

575

with the Bead-C3Sensor + caspase 3 incubation shows a detectable difference in reporter

576

fluorescence emission channel.

is applied to the Bead-C3Sensor after 3 hour incubation with active Caspase-3 protein. On

577



578 579

Figure 4. Small molecule identification tag, design and amplification. (a) Design of barcode

580

component. (b) Electrophoresis DNA gel after PCR amplification of the barcode. The

581

photocleavable linker was necessary for adequate amplification. Using similar thermocycler

582

conditions and identical primers, barcodes of different sequences were amplified with similar

583

efficiency. There is no bias observed for one sequence over the other. (c) Electrophoresis DNA

584

gel after PCR amplification of the barcodes incubated with cell lysate. A band at the appropriate

585

molecular weight marker is observed, demonstrating that the oligonucleotide sequence is

586

maintained throughout lysis conditions and exposure to enzymes and cellular components

587

present during cell death by apoptosis.

588 589

Figure 5. Bead-FITC-Barcode internalization by HT-1080 cells. (a) Confocal microscopy

590

internal slice of Z-stack image, after Bead-FITC-Barcode technology incubation with HT-1080

591

cells for 6 hours. (i) Bead fluorescence in red bandpass filter. (ii) FITC fluorescence of FITC-

592

peptide conjugated to bead in green bandpass filter. (iii) Transmitted light. (iv) DAPI (v) Overlay.

593

(vi) Full Z-stack overlay image. (b) Slices of Z-stack produced by confocal microscopy of Bead-

594

FITC-Barcode technology with HT-1080 incubation. Yellow overlay shows co-localization of

595

Bead-FITC signal. Outermost slices show no Bead presence, supporting the observation that

596

the cell internalizes the bead. (c) HT-1080 cells incubated for 72 h with varying concentrations

597

of bead conjugated to the barcode and fluorescent reporter show no decrease in cell viability.

598 599

Figure 6. Small molecule component design and attachment to bead. (a) Structure of

600

Bead-Disulfide linker-DM1. (b) Structure of Bead-Malemide linker-DM1. (c) Structure of Bead-

601

Cathepsin D cleavable linker – monomethyl Auristatin E (MMAE). (d) Neither thiol-reduction

602

based-cleavable nor non-cleavable linkers conjugated to the beads show lethality when

603

incubated with HT-1080 cells in varying concentrations. Cathepsin D cleavable linker- based


Page 30 of 34

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604

bead system shows lethality, demonstrating effective release of MMAE upon cellular

605

internalization.

606 607

Acknowledgements

608

C. Yozwiak was funded by the NCI Training Program in Cancer Biology Grant T32GG007284-

609

13. This research of B. R. Stockwell was funded by NIH/NCI (1R35CA209896 and

610

P01CA087497).

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Supporting Information Supporting information is available online, including Supplementary Figure S1 to S6 and Tables S1 and S2, which show the synthetic scheme to create the delivery system, uptake analysis of beads, characterization of the C3 sensor, size analysis of beads, testing of the disulfide linker, and small molecule controls, and the peptide and barcode sequences ordered. his material is available free of charge via the internet at http://pubs.acs.org.



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References

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