
==== Front
Hepat Oncol
Hepat Oncol
Hepatic Oncology
2045-0923
2045-0931
Taylor & Francis

10.1080/20450923.2024.2389031
2389031
Version of Record
Research Article
Research Article
APT imaging of hepatocellular carcinoma signals an effective therapeutic response in advance of tumor shrinkage
Wang Xiaojing a
Ishimatsu Keisuke a
Li Junjie a
Wen Xiaodong a
Ou Weijun a b
Anwar Arnida a
Chaudhary Jaideep a
Takahashi Masaya a
Sherry A Dean a c
https://orcid.org/0000-0001-6902-8034
Corbin Ian R * a c d
a Advanced Imaging Research Center, University of Texas Southwestern Medical Center at Dallas, Dallas, TX 75390, USA
b Organ Transplantation Center, The First Affiliated Hospital, Sun Yat-sen University, Guangzhou, PR China
c Internal Medicine Division of Liver & Digestive Diseases, University of Texas Southwestern Medical Center at Dallas, Dallas, TX 75390, USA
d Department of Radiology, University of Texas Southwestern Medical Center at Dallas, Dallas, TX 75390, USA
* CONTACT: Tel.: +1 214 645 7044; ian.corbin@utsouthwestern.edu
12 9 2024
2024
12 9 2024
11 1 2389031Aptara05 8 2024
10 9 2024
15 8 2023
23 7 2024
© 2024 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (http://creativecommons.org/licenses/by-nc/4.0/), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited. The terms on which this article has been published allow the posting of the Accepted Manuscript in a repository by the author(s) or with their consent.

Aim: The aim of this study was to assess the utility of weighted amide proton transfer (APTw) MRI in three different rodent models of hepatocellular carcinoma (HCC).

Methods: APTw MRI was evaluated in models of diethylnitrosamine (DEN) induced HCC, N1S1 syngeneic orthotopic xenograft and human HepG2 ectopic xenograft.

Results: All models of HCC showed a higher APTw signal over the surrounding normal tissues. In the DEN model, the APTw signal could differentiate HCC lesions from benign nodules. Intra-arterial administration of low-density lipoprotein docosahexaenoic acid (LDL-DHA) nanoparticles to N1S1 xenografts rapidly lowered the tumor APTw signal within 72 h. Direct injections of LDL-DHA nanoparticles into HepG2 xenografts also showed similar therapeutic responses.

Conclusion: We have demonstrated the utility of APTw imaging in the diagnostic/therapeutic management of HCC.

Plain Language Summary

The aim of this study was to evaluate a specific imaging method called amide proton transfer (APT) MRI in different animal models of primary liver cancer. Three models of liver cancer were developed by chemical induction (in rat) and transplantation of rat and human tumor cells in rat and mouse respectively. A higher APTw signal was detected in all tumor models of liver cancer. In the chemical induction model, APTw MRI could even distinguish between cancer and benign growths in the liver. Treatments with anticancer nanomedicines rapidly decreased tumor APTw signals which was accompanied with significant tumor cell death. These findings demonstrated that APTw imaging is a useful tool in the management of liver cancer.

Tweetable Abstract

This study examined the utility of APT MRI in three animal models of liver cancer. Data revealed that APTw MRI can identify liver tumors from normal liver and can detect tumor killing following drug treatment. ATP MRI may be a useful tool for managing subjects with liver cancer.

Article highlights

Background & aim

The MRI APTw signal is an innovative molecular imaging biomarker that is significantly enhanced in malignant tissues relative to normal tissues.

Three preclinical animal models of HCC where employed in this study to evaluate the utility of APTw imaging in HCC.

ATP imaging for HCC detection

Chemical induced HCC (DEN), syngeneic orthotopic xenograft (N1S1 hepatoma) and ectopic human xenografts (HepG2) all displayed and increased APTw signal over their surrounding normal tissues.

APTw imaging was able to differentiate HCC lesions from dysplastic/regenerative nodules.

ATPw imaging for monitoring response to therapy

Intra-arterial locoregional administration of LDL-DHA nanoparticles to N1S1 orthotopic xenografts selectively showed a progressive decrease in tumor APTw signal over 72 h post injection.

The loss of APTw signal in the N1S1 tumor correlated with treatment induced tumor necrosis.

Direct injections of LDL-DHA nanoparticles into ectopic human HepG2 xenografts similarly showed depletion of the tumor APTw signal and pronounced tumor destruction.

Summary

APTw imaging proved to be highly sensitive to LDL-DHA mediated treatment of HCC.

APTw imaging may prove to be a valuable tool in the diagnostic and therapeutic management of HCC.

Keywords: 

docosahexaenoic acid
hepatocellular carcinoma
low-density lipoprotein
molecular imaging
nanoparticle
National Cancer Institute 10.13039/100000054 R01CA215702 Remeditex Ventures LCC OTD-109946 This work was supported in part by NCI, National Institutes of Health (NIH), Grant R01CA215702; Remeditex ventures LCC OTD-109946; the UTSW Cancer Center Support Grant (5P30 CA 142543-05) and UTSW Center for Translational Medicine Grant (UL1TR001105).
==== Body
pmc1. Introduction

Hepatocellular carcinoma (HCC) is the most common primary liver cancer and the fourth leading cause of cancer-related deaths worldwide [1]. Early detection of HCC is vital to enabling potentially curative treatments such as hepatic resection or liver transplantation. As such, noninvasive radiologic imaging plays a critical role in the management of HCC [2]. Unlike most solid cancers, the diagnosis and treatment of HCC can be rendered, based on noninvasive imaging without histopathology confirmation. Contrast enhanced multiphase magnetic resonance imaging (MRI) or computed tomography (CT) are typically employed to diagnose HCC based on features of ‘arterial enhancement and delayed washout’, in other words, hyper-vascularity during the arterial phase and low signal intensity on the portal venous or delayed phases of imaging. This classic appearance is attributed to the liver's dual blood supply, where the normal healthy liver receives most of its blood supply from the portal vein and HCC lesions primarily obtain their blood supply from hepatic artery branches. This enhancement pattern provides a specificity of 95% for the diagnosis of HCC [3,4]. Beyond HCC detection, noninvasive imaging also plays an essential role in assessing tumor response to treatment [2]. For patients with early and intermediate staged HCC who receive local ablative and transarterial locoregional treatments, contrast enhanced MRI and CT provides multiparametric assessments of tumor viability. Assessment algorithms such as the modified Response Evaluation Criteria in Solid Tumors (mRECIST), which use change in lesion size/number, lesion characteristics and viable portions of the lesions as determined by arterial phase enhancement to define the response to treatment [5]. The mRECIST criteria categorize treatment response for HCC lesions as complete, partial, stable disease, progressive disease, or development of new lesion(s) [5].

As described above, contrast enhanced CT and MRI play a central role in the management of HCC with good overall sensitivity and specificity. However, only HCCs that have sufficient neoangiogenesis show arterial hyper-enhancement and exhibit contrast agent washout can be unequivocally diagnosed. Up to approximately 40% of HCCs lack arterial phase hyper-enhancement [6,7] and cannot be diagnosed as definite HCC using contrast enhanced imaging. These include most early HCCs [8]; poorly differentiated, infiltrative HCCs [9]; HCCs with foci of hyper-vascularity [6]; and hyper-enhancing HCCs missed due to arterial phase mistiming or imaging artifacts. Moreover, an estimated 40%–60% of small HCCs, even if displaying arterial phase hyper-enhancement, do not show a washout or capsule appearance in the venous phases [10,11], and so cannot be diagnosed as definite HCC using only contrast agents. Collectively, these concerns highlight the need for non-contrast enhanced imaging approaches for HCC management.

In recent years, amide proton transfer (APT) imaging has been introduced as a novel contrast mechanism in the field of molecular imaging [12]. The technique is performed without injection of an exogenous contrast agent and is based on chemical exchange saturation transfer (CEST) of labile protons in biomolecules with solvent water protons [13]. APT is a subset of CEST that refers specifically to exchange of amide protons resident on the backbone of low concentration mobile cytosolic proteins and peptides [14]. Signal enhancement factors of up to 106 have been reported for some systems [15]. Rapid proliferation and high cell density characteristic of malignant tissues is thought to be accompanied by elevated levels of cytosolic protein and, in turn, an increased APT signal [14,16]. APT imaging has previously been used to identify [17–19], grade [20,21] and differentiate radiation necrosis from recurrent tumor growth [22,23] in various tumor types. In addition, in recent years there has been increased appreciation for the contributions of intracellular tissue pH to the APT signal. Seminal studies performed by Ray et al. estimate that 66% and 34% of the APT signal in tumors arise from changes in cytosolic protein and pH, respectively [24]. The goal of the present study is to assess the utility of APT imaging to detect malignant tumors in preclinical rodent models of HCC. Secondly, we aim to evaluate the ability of APT imaging to monitor the treatment response of HCC bearing rodent models to a novel lipoprotein-docosahexaenoic acid-based nanomedicine called LDL-DHA.

2. Materials & methods

2.1. Preparation of LDL-DHA nanoparticles

Human LDL was isolated from apheresis plasma of patients with familial hypercholesterolemia using sequential density gradient ultracentrifugation. Incorporation of unesterified DHA (Nu-chek Prep, Inc, MN, USA) into LDL was performed by the reconstitution method, as described in our previous publication [25]. In these studies, LDL reconstituted with oleic acid (LDL-OA) or triolein (LDL-TO) served as control nanoparticles. Nanoparticle characterization (structure and composition) was performed as described previously to ensure consistency of batch to batch preparations.

2.2. Cell culture

N1S1 rat hepatoma cell line (ATCC, CRL-1603, VA, USA) and the human liver tumor cell line HepG2 were cultured in Dulbecco's Modified Eagle's Medium (Sigma, D6429) supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin. Cells were incubated at 37°C in a humidified environment containing 5% CO2.

2.3. Animal studies

All animals were kept at a temperature controlled (22 ± 2°C) room with a light-dark cycle of 12 h and fed with standard laboratory food and tap water ad libitum throughout the studies in accordance with the National Research Council (US) Committee Guide and Care and Use of Laboratory animals. All animal studies were approved and conducted under the oversight of the UT Southwestern Institutional Animal Care and Use Committee. NIH OLAW Assurance Number D16-00296.

2.4. Diethylnitrosamine (DEN) HCC model

Autochthonous HCCs was induced in 4 week old male Wistar rats (Charles River Laboratories) using ad libitum oral intake of 0.01% DEN (Sigma-Aldrich, St. Louis, Missouri) in their drinking water for 12 weeks. The water bottles were changed twice per week with freshly prepared 0.01% DEN. At the end of the 12 week exposure to DEN, nearly all animals presented with multiple HCCs on a background of severe liver fibrosis/cirrhosis [26,27]. After the 12 week induction period, animals were returned to regular water for an additional 2 weeks (to allow recovery from acute hepatitis) before imaging.

2.5. Xenograft models

2.5.1. Orthotopic xenograft studies

A syngeneic orthotopic rat hepatoma model was established in 7 week old male Sprague-Dawley rats (Envigo) using the rat N1S1 heptoma cell line. Rats were anesthetized with isoflurane (2.5%) and the abdomen of each rat was shaved and prepared for aseptic surgery, a 2 cm vertical midline laparotomy was performed, and the left liver lobe was exposed. N1S1 cells (7 × 106) in 50 μl of phosphate-buffered saline solution mixed with Matrigel (Corning Life Sciences, Corning, NY) at 1:1 ratio were injected into the left upper liver lobe. The abdominal cavity was then closed in two layers with 3.0 silk sutures and analgesia was administrated every 10 hrs for 3 days. All experiments were initiated ∼12 days after cell implantation.

2.5.2. Intrahepatic arterial cannulation

Tumor-bearing rats were allocated to group 1 or 2. Group 1 rats received an intrahepatic arterial injection of LDL-DHA nanoparticles (2.0 mg/kg); group 2 (control group) received injection of LDL reconstituted with oleic acid (LDL-OA, 2.0 mg/kg). Locoregional access to the hepatic artery was performed. The gastroduodenal artery of tumor-bearing rats was isolated with two 6.0 sutures, and the tip of a 2F silicone microcatheter (Access Technologies, Skokie, Ill) was advanced through an arterial incision made between the two sutures to the proper hepatic artery, and the artery was ligated with a slipknot from both sides to fix the catheter in place. Injections were given at a rate of 6 ml per hour. The catheter was then carefully withdrawn from the arterial incision and the suture was gently tightened to prevent bleeding. The gastroduodenal artery was then permanently ligated, and the abdominal cavity was sutured in two layers.

2.5.3. Ectopic xenograft studies

The ectopic HCC xenografts were prepared as previously described [28], briefly male Balb/C severe combined immunodeficiency (SCID) mice (6–8 weeks old; n = 5 per group) were obtained from Charles River Laboratory. HepG2 cells (5 × 106 cells) suspended in phosphate-buffered saline (PBS) were injected subcutaneously into the right flank of mice. On day 15 after implantation of tumor cells and the tumors were about 3.0–5.0 mm in diameter, animals were randomly allocated to receive either LDL-DHA or control LDL-TO control nanoparticle treatments. Pre-treatment APTw imaging was performed at this time. Mice were anesthetized and intratumoral injections were directed toward the center of the tumor. The needle was inserted once into the predefined region, and 100 μl of the LDL nanoparticle solution was dispensed with even pressure. The injection needle was removed slowly, and the injection site was sterilized. LDL nanoparticles were administered at an equivalent dose of 2.0 mg/kg LDL-DHA. Mice received single intratumoral injections for two consecutive days followed by 1 day of recovery. On the fourth day, all mice were imaged, sacrificed and the tumors removed for volume measurements. The largest and smallest diameters were measured with vernier calipers and tumor volume was estimated according the formula: V = 1/2 ab2, where a and b are the largest and smallest tumor diameters, respectively, and V is the tumor volume in milliliters. After tumor volume measurements, each tumor was divided, fixed in formalin and prepared for histology.

2.5.4. APTw imaging

Tumor-bearing rats were imaged using an Agilent (Varian) 9.4T small-animal MR system (equipped with a 72-mm (I.D.) volume coil) to monitor tumor growth and access tumor response to LDL-DHA nanoparticle treatment or LDL-OA/LDL-TO control. Axial multi-slice T2-weighted images were collected covering the entire liver using a fast spin-echo sequence. Upon choosing a single 2-mm slice that showed maximum tumor area, APTw imaging was performed under respiratory gating. The MRI data were obtained using a steady state gradient echo pulse sequence (GEMS) with centric K-space encoding preceded by a rectangular saturation pulse (B1, 2.3 μT; duration, 5 s) applied at 29 frequency offsets from 7 to -7 ppm with an interval of 0.5 ppm. A control image with the saturation offset at 300 ppm was also acquired. The body temperature was regulated at 37.0 ± 0.1 °C with an animal monitoring system from Small Animal Instruments (Stony Brook, NY).

2.5.5. MR data analysis

All image data were analyzed using self-written scripts in MATLAB R2018a (The Mathworks, Inc., Natick, MA, USA) and ImageJ (version 1.48v, National Institutes of Health, Bethesda, MD). The magnetization transfer ratio (MTR) is defined as: MTR = 1 - Ssat/S0, where Ssat and S0 are the signal intensities with and without presaturation pulse, respectively. In the data processing, the images obtained at 29 frequency offsets were first organized to generate a z-spectrum (Supplementary Figure S1). The z-spectrum was corrected pixel-wise for B0 inhomogeneity effects by interpolation and centering of the water protons signal to zero ppm. The B0 inhomogeneity was measured using the Water Saturation Shift Referencing (WASSR) method. A total of 31 images using a 0.2 μT presaturation pulse of 2 s duration collected over ± 1.5 ppm steps of 0.1 ppm were used for B0 correction. A MTR asymmetry (MTRasym) map is then generated by subtracting [Ssat(+offset)/S0]–[Ssat(-offset)/S0] in each pixel. A MTRasym map generated after applying a pre-saturation pulse at ± 3.5 ppm is referred to as an APT-weighted image (APTw). These are referred to as APTw because there are other protons in the +3.5 ppm chemical shift region in exchange with bulk water protons so the net signal cannot be attributed to amide exchanging protons only. Furthermore, the bulk water proton signal intensity can also be altered after a presaturation pulse is applied at -3.5 ppm due to upfield NOE signals [29], so the APTw image can, in principle, be either negative (if the NOE signal is high) or positive (if the amide proton signal is high). In normal health liver tissue, the NOE signal at -3.5 ppm is typically higher than amide proton exchange signal at +3.5 ppm so the difference appears as a net negative APTw signal. In tumors, the amide proton exchange signal typically increases and becomes higher than the NOE signal so, in this case, the APTw signal is positive. ROIs were either drawn over the entire tumor or in smaller regions within each tumor to evaluate APTw for that region. ROIs were also drawn in adjacent normal liver and used for reference.

2.5.6. Histopathological analysis

At the time of euthanasia, excised liver and tumor samples were collected, and were fixed in 10% neutral buffered formalin for 24h, then embedded in paraffin, and sectioned into 5-μm slices and stained with Hematoxylin and Eosin (H&E). The slides were captured with an optical microscope (CX31, Olympus, Japan) at 100× magnification for histopathology, and micrograph were taken with a microscope digital camera system (DP50, Olympus, Japan).

2.5.7. Data availability

The data generated in this study are available within the article and its Supplementary data files or are available upon request from the corresponding author.

3. Results

3.1. HCC detection in DEN induced HCC

Serum liver function test results for untreated controls and DEN-treated rats are presented in Table 1. Like earlier reports, DEN treated rats had significantly higher levels of serum ALT, AST, ALKP, GGT and TBIL. These findings demonstrate hepatic dysfunction indicative of chronic liver disease.

Table 1. Serum biochemistry of Control and rats exposed to 12 weeks of DEN.

Serum parameters	Control	DEN	
ALT(U/l)	50.7 ± 0.6a	170.6 ± 35.7b	
AST(U/l)	65.8 ± 4.3a	275.8 ± 65.8b	
GGT(U/l)	5.0 ± 0.0a	27.2 ± 9.0b	
TBIL(mg/dl)	0.5 ± 0.1a	0.7 ± 0.1a	
ALB(g/dl)	3.1 ± 0.1a	2.7 ± 0.2a	
ALKP(U/l)	96.8 ± 8.1a	151.0 ± 12.9b	
a or b Means with different letters are significantly different from each other, p ≤ 0.05.

Data are expressed as mean ± SEM.

ALB: Albumin; ALKP: Alkaline phosphatase; ALT: Alanine aminotransferase; AST: Aspartate aminotransferase; GGT: Gamma glutamyltransferase; TBIL: Total bilirubin.

After completion of 12 weeks of DEN exposure, all rats displayed multiple liver lesions (Figure 1A & B). The irregular contour of the liver on T2W images is indicative of liver cirrhosis. Focal liver lesions typically appeared hyperintense in T2W images (Figure 1A). Histological findings confirmed altered hepatic architecture consistent with liver cirrhosis and foci of malignant tumors (Figure 1C & D). Representative anatomical and ATPW images of DEN treated rats are shown in Figure 2. This representative APTw image slice was selected to illustrate the maximum signal in the liver and tumor. Other slices through the liver/tumor displayed similar APTw images. These results showed that gated APTw imaging is feasible in the free breathing rat. It should be noted that APTw imaging detects only those protons in exchange with water in this chemical shift region (3.5 ppm downfield of water), not the entire pool of water protons. As such the APTw signal is typically only ∼1–6% the intensity of bulk water protons and this yields in a much lower resolution image compared with a standard T1w or T2w image. For this reason, the APTw image shown in Figure 2 appears to have a slightly different shape than the anatomical T2w image because the resolution of the two images differ considerably. Quantitatively, the measured APTW signal detected in liver lesions (28 hepatic lesions were identified in 16 rats) ranged in intensity from -1.6 to 6.2 while APTW signal of the surrounding normal liver ranged in intensity from -4.5 to -0.3. Lesions with an APTW signal intensity >0 were histologically confirmed to be malignant HCC. Lesions in the -1.5 to -0.3 intensity range displayed features consistent with benign dysplastic or regenerative nodule (Figure 3). It should be noted that APTW values for the surrounding liver also approached readings in the -1.5 to -0.3 range, which is expected given the regenerative and premalignant state of the DEN treated liver [30]. It stands to reason that APTw values between -0.3 and 0 may reflect liver tissue in the dedifferentiation continuum from dysplastic nodules to early-stage malignant HCC. The present findings demonstrate that APT imaging is a sensitive radiologic method for identifying malignant hepatic lesions in the setting of liver fibrosis.

Figure 1. Imaging, photography and histology of rat following 12 weeks of diethylnistrosamine (DEN) induced HCC. (A) Axial T2W MRI through the abdomen of DEN treated rat. Note the hyper-intense lesions in the liver. CEST phantom (25 mM arginine). (B) Photograph of excised liver bearing DEN induced HCC and liver nodules. (C) liver histology from DEN treated rat. (D) Histology of HCC from DEN treated rat. Images captured at 20x magnification. Scale bar = 100 um.

Figure 2. Representative APT imaging of rat with diethylnistrosamine (DEN) induced HCC. (A) Axial T2- weight image. (B) APT imaged co-registered with T2 axial MRI. The ATP image masks are applied to liver, tumor and spinal muscle. (C) scatter plot of APT readings from liver parenchyma and tumor regions of DEN rats.

**** represents p < 0.0001 difference between groups.

Figure 3. Paired scatter plot showing corresponding APTw values of liver parenchyma and hepatocellular lesion from the same individual animal. (A) Data for entire cohort; (B) data from malignant HCC lesions; and (C) data from benign regenerative nodule. Scale bar = 100um.

3.2. Assessment of transarterial locoregional LDL-DHA treatment in a syngeneic orthotopic model of HCC

N1S1 tumors were visible in liver by T2W MRI 5 days after cell inoculation. Histology confirmed the tumors were malignant HCC. APT imaging was then performed before and after a single dose of LDL-DHA nanoparticles or LDL-OA control particles (2.0 mg/kg) to evaluate tumor necrosis and response to treatment. The average size (maximum diameter) of all tumors was 12.9 ± 1.1 mm (n = 8). APT imaging was performed under respiratory gating with data collection triggered at end respiratory phase following a presaturation pulse.

Figure 4 shows representative APTW images of a N1S1 liver tumor prior to treatment. Prior to treatment with LDL-OA control nanoparticles, there was a clear increase in the APTw signal intensity in the tumor mass (close arrow), compared with the surrounding normal-appearing liver (open arrow). In this LDL-OA treated rat, the size (maximum diameter) of the tumor was increased from 14.7 mm to 15.2 mm at 1 day post-treatment, and continuously increased to 16.6 mm after 3 days post-treatment. Similarly, the APTW signal did not change significantly over the first 24 h period (-1.6 ± 0.3% to -0.1 ± 0.6%, 1 day post-treatment) but did increase to -0.5 ± 0.4% by 3 days post-treatment (Figure 4 upper insert). This indicates continued growth of the tumor over this 3 day period. Histology of the excised tumors confirmed the viability of the LDL control treated tumor (Figure 4 lower insert).

Figure 4. Changes in APT signal intensity (as a percentage) for the N1S1 tumors following in LDL-OA nanoparticle treatment. (Upper panel) T2-weighted axial MRI (lower panel) APT imaged co-registered with T2 axial MRI. The ATP image masks are overlaid the liver and tumor region. (Insert upper) Change in the APT signal in liver and tumor tissue before, 24 and 72 h post LDL-OA treatment. (Insert lower) H&E stained section showing liver and tumor interface. White arrows indicate tumor. Scale bar = 100 um.

Figure 5 shows the representative APTW images of N1S1 tumors in the LDL-DHA nanoparticle-treated group. In this case, the size (maximum diameter) of the tumor also increased from 17.6 mm to 20.7 mm at 1 day post-treatment, but then decreased in size to 19.9 mm at 3 days post-treatment. Unlike the tumor mass, the LDL-DHA treated tumors displayed a pronounced drop in APTW signal, from -1.9 ± 1.4% to -3.7 ± 1.8% at 1 day post-treatment, and an even larger decrease at 3 days post-treatment (-6.8 ± 1.6%). Histology of the excised tumors confirmed the extensive tumor cell necrosis induced by the LDL-DHA treatment (Supplementary Figure S2).

Figure 5. Changes in APT signal intensity (as a percentage) for the N1S1 tumors following in LDL-DHA nanoparticle treatment. (Upper panel) T2-weighted axial MRI (lower panel) APT imaged co-registered with T2 axial MRI. The ATP image masks are overlaid the liver and tumor region. (Insert upper) Change in the APT signal in liver and tumor tissue before, 24 and 72 h post LDL-DHA treatment. (Insert lower) H&E stained section showing liver and tumor interface. White arrows indicate tumor.

*** represents p < 0.001 difference between groups. Scale bar = 100 um.

3.3. Monitoring LDL-DHA Treatment in Ectopic Human HCC Xenograft

APTw imaging was also performed in HepG2 xenograft bearing-mice before and after two local treatments of either LDL control nanoparticles (n = 6, labeled LDL-TO in Figure 6) or LDL-DHA nanoparticles (n = 6, labeled LDL-DHA in Figure 6). In the LDL control treated tumors, the APTW signal increased after treatment indicating growth of viable tumor tissue. Conversely, the LDL-DHA treated tumors displayed a pronounced drop in APTW signal, indicative of a significant treatment effect. On average, the APTW signals in the LDL-DHA treated tumors were 70% lower than that detected in the LDL control treated tumors (Figure 6). Histopathology of the excised tumors confirmed the viability of the LDL control treated tumors and the extensive tumor cell necrosis induced by the LDL-DHA treatment (Figure 7).

Figure 6. APT weighted axial images of mice with HepG2 (HCC) xenografts before and after local LDL nanoparticle treatment. (A) Tumor volume and APT signal increase after LDL control (LDL-TO) treatment while APT signal is significantly reduced after LDL-DHA treatment. White arrow indicates tumor; red sphere, CEST phantom (25 mM arginine); APT signal from dorsal muscle also displayed. (B) The average APT signal was 2.3 ± 0.5 vs 0.7 ± 0.4 for LDL-TO and LDL-DHA treated tumors respectively (n = 6 mice/group).

**p < 0.05.

Figure 7. LDL-DHA treatments inhibit HCC tumor growth. Photograph of HepG2 tumor following intratumoral injection of LDL-TO control (A) or LDL-DHA (B) treatments. Representative excised tumors are displayed below. (C) Tumor histology following LDL-TO treatments. (D) Tumor histology following LDL-DHA treatments. Scale bar = 100 um.

4. Discussion

In the present study, we evaluated the diagnostic utility of APTW imaging in three preclinical animal models of HCC. In each model, whether chemically induced, orthotopic or ectopic xenograft, the HCC tumors consistently showed a higher APTW signal over the surrounding normal tissues. This has previously been reported for high-grade HCC [21,31] and has consistently been observed across numerous malignancies [14,16,32,33]. The APTW signal has contributions from multiple sources including possible alterations in tissue amide proton concentration, changes in tissue pH and NOE contributions. The average chemical shifts of typical backbone amide protons in proteins and peptides appear between 8.2–8.4 ppm (3.5–3.7 ppm downfield of water) [34], so it is reasonable to assume that the APTW signals detected in HCC tumors have at least some contribution from changes in protein concentration (i.e. cellular density and proliferation rate). Indeed, rapidly proliferating cells have increased rates of protein synthesis [35] and several publications have suggested that certain tumor types have increased protein relative to their normal tissue [14,16]. However, other biological studies support that the concept that intracellular protein levels are proportional to cell volume and thus protein concentrations remain constant throughout the cell cycle [36]. Other factors beyond absolute protein concentration may also contribute to changes in the APTW signal. The number of amide protons available for exchange per protein molecule (i.e. the chemical basis of CEST) could reflect differences in protein conformation, proteolysis rates or protein size. Each of these processes are expected to occur during rapid cell proliferation.

Given that the exchange rate of amide protons with solvent water protons is a base catalyzed reaction, the APTW signal is also quite sensitive to pH. HCC like other malignant tissues have aberrant metabolism that results in extracellular acidification [37] with a concomitant increase in intracellular pH [38,39]. Although such increases in intracellular in pH may be small, this could contribute to an increase in tumor APTW signal as the tumor grows because -NH proton exchange is exquisitely sensitive to pH above pH ∼7.4. For example, a recent imaging study reported that 66% of the measured APTW signal in brain tumors arose from tissue protein content while 34% of the signal was ascribed to an increase in intracellular pH [24].

The chemically induced DEN HCC model was examined first in this study. This model is commonly used in preclinical studies as it recapitulates many of the clinical features seen in HCC patients which include background liver dysfunction, hepatic fibrosis and focal benign and malignant lesions [27,40]. The latter arises as a result of DEN induced oncogenic mutations that drive the progressive formation of regenerative nodules, premalignant dysplastic lesions and overt malignant HCC [41]. After 12 weeks of DEN treatment, this range of nodular lesions can be seen along with established liver fibrosis and signs of hepatic impairment. T2-weighted MRI revealed numerous focal hepatic lesions in DEN treated rats which appear hyperintense compared with the normal liver. This finding, however, provides little in the way of differential diagnosis as regenerative nodules, dysplastic lesions and overt HCC have all been shown to display variable signal intensity characteristics in T2w images [42]. Subsequent APTw imaging revealed that these T2W lesions could be further distinguished as having high, moderate or low APTw signals. The focal lesions with high APT signal intensity were confirmed by histopathology to display a disorganized architecture, increased cellular density and enlarged hyperchromatic nuclei consistent with malignant HCC phenotype while the lesions with low APTw signal intensity were identified as regenerative or dysplastic nodules. Collectively, these findings suggest that APTw imaging can provide an additional level of differentiation over T2W images in distinguishing malignant from benign hepatic lesions.

The role of APTw imaging in evaluating the response to therapy has been documented by numerous investigators [43–45]. Tumors that respond to chemotherapy or radiation typically display a reduced APTw signal intensity by 6–7 days post treatment [43,45]. This timeline is indicative of the temporal induction of tumor cell death (necrosis/apoptosis) initiated by these therapies. In the current study, the therapeutic effects of the novel nanomedicine, LDL-DHA, was investigated in rodent xenograft models of cancer. Previous studies have demonstrated that LDL-DHA elicits lethal accumulation of lipid ROS and depletion of cellular glutathione [28,46]. Following locoregional transarterial administration of LDL-DHA into syngeneic HCC-bearing rats, a precipitous drop in tumor APTw signal was seen as early as 1 and 3 days post treatment. The fast response detected by APTw imaging reflects the potent and rapid metabolic perturbations induced by the LDL-DHA therapy. Furthermore, the treatment also demonstrated high tumor selectivity as the APTw signal from healthy tissue regions of livers of treated animals remained unaltered. Histopathological examinations confirmed the imaging findings as the HCC displayed near complete necrosis, while the surrounding liver tissue remained largely unaffected. The ectopic human HCC mouse model was employed to demonstrate the utility of this therapy against human malignancy. Similar to the rat HCC model, the human HCC (HepG2) cells were equally responsive to direct LDL-DHA treatment. In HepG2 tumors, the APTw signals plummeted after LDL-DHA treatment compared with control treated tumors. The corresponding histopathology also showed massive necrosis in LDL-DHA treated tumors, while viable tissue was evident throughout control treated tumors.

Collectively, these findings indicate that APTw imaging is highly sensitive to selective tumor necrosis initiated by LDL-DHA treatment. LDL-DHA creates a highly oxidative tumor microenvironment replete with lipid peroxides and, under these conditions, protein conformations, proteolysis rates and amide proton exchange/pH are certain to change. Compounded with the loss of cellular integrity, the contribution of mobile protein/peptide -NH proton exchange to the APTw images is reduced, perhaps due to loss of cytoplasm.

5. Conclusion

In conclusion, this study demonstrated that APTw imaging is a useful imaging biomarker to detect malignant versus benign lesions in the liver. Furthermore, the APTw signal is also sensitive to treatment responses following liver-directed LDL-DHA therapy. Such technology could potentially impact the management of patients with liver tumors.

Supplementary Material

Supplementary Materials

Supplemental material

Supplemental data for this article can be accessed at https://doi.org/10.1080/20450923.2024.2389031

Author contributions

X Wang: Acquisition and analysis of data; study design; draft article. K Ishimatsu: Acquisition and analysis of data; study design. J Li: Acquisition and analysis of data; study design. X Wen: Acquisition and analysis of data. W Ou: Acquisition and analysis of data; study design. A Anwar: Acquisition of data. J Chaudhary: Acquisition and analysis of data; draft article; critical revision of content. M Takahashi: Advice and support for the study. AD Sherry: Advice and support for the study, editing of manuscript. IR Corbin: Contribution to conception and design; analysis and interpretation of data; draft article and critical revision; final approval for publication; accountable for all aspects of work.

Financial disclosure

This work was supported in part by NCI, National Institutes of Health (NIH), Grant R01CA215702; Remeditex ventures LCC OTD-109946; the UTSW Cancer Center Support Grant (5P30 CA 142543-05) and UTSW Center for Translational Medicine Grant (UL1TR001105). The authors have no other relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript apart from those disclosed. Permission is granted to deposit the present article in the NIHMS system/PMC.

Competing interests disclosure

The authors have no competing interests or relevant affiliations with any organization or entity with the subject matter or materials discussed in the manuscript. This includes employment, consultancies, honoraria, stock ownership or options, expert testimony, grants or patents received or pending, or royalties.

Writing disclosure

No writing assistance was utilized in the production of this manuscript.

Ethical conduct of research

UT Southwestern uses the “Guide for the Care and Use of Laboratory Animals” when establishing animal research standards. All animal studies were approved and conducted under the oversight of the UT Southwestern Institutional Animal Care and Use Committee.

Data availability statement

All data generated or analyzed during this study are included in this article and its Supplementary material files. Further enquiries can be directed to the corresponding author.
==== Refs
References

Papers of special note have been highlighted as: • of interest

1. Bray F, Ferlay J, Soerjomataram I, et al. Global cancer statistics 2018: GLOBOCAN estimates of incidence and mortality worldwide for 36 cancers in 185 countries. CA Cancer J Clin. 2018;68 (6 ):394–424. doi:10.3322/caac.21492 30207593
2. Marrero JA, Kulik LM, Sirlin CB, et al. Diagnosis, staging, and management of hepatocellular carcinoma: 2018 Practice Guidance by the American Association for the Study of Liver Diseases. Hepatology. 2018;68 (2 ):723–750. doi:10.1002/hep.29913 29624699
3. Marrero JA, Hussain HK, Nghiem HV, et al. Improving the prediction of hepatocellular carcinoma in cirrhotic patients with an arterially-enhancing liver mass. Liver Transplantation. 2005;11 (3 ):281–289. doi:10.1002/lt.20357 15719410
4. Tang A, Bashir MR, Corwin MT, et al. Evidence supporting LI-RADS major features for CT- and MR imaging–based diagnosis of hepatocellular carcinoma: a systematic review. Radiology. 2018;286 (1 ):29–48. doi:10.1148/radiol.2017170554 29166245
5. Lencioni R, Llovet JM. Modified RECIST (mRECIST) assessment for hepatocellular carcinoma. Semin Liver Dis. 2010;30 (1 ):52–60. doi:10.1055/s-0030-1247132 20175033
• Describes standard sets of criteria used to evaluate the tumor response of hepatocellular carcinoma (HCC) treated using loco-regional therapy.

6. Sano K, Ichikawa T, Motosugi U, et al. Imaging study of early hepatocellular carcinoma: usefulness of gadoxetic acid–enhanced MR imaging. Radiology. 2011;261 (3 ):834–844. doi:10.1148/radiol.11101840 21998047
7. Yoon SH, Lee JM, So YH, et al. Multiphasic MDCT enhancement pattern of hepatocellular carcinoma smaller than 3 cm in diameter: tumor size and cellular differentiation. AJR Am J Roentgenol. 2009;193 (6 ):W482–489. doi:10.2214/AJR.08.1818 19933622
8. Kim SH, Lee WJ, Lim HK, et al. SPIO-enhanced MRI findings of well-differentiated hepatocellular carcinomas: correlation with MDCT findings. kjr. 2009;10 (2 ):112–120. doi:10.3348/kjr.2009.10.2.112
9. Kanematsu M, Semelka RC, Leonardou P, et al. Hepatocellular carcinoma of diffuse type: MR imaging findings and clinical manifestations. J Magn Reson Imaging. 2003;18 (2 ):189–195. doi:10.1002/jmri.10336 12884331
10. Sangiovanni A, Manini MA, Iavarone M, et al. The diagnostic and economic impact of contrast imaging techniques in the diagnosis of small hepatocellular carcinoma in cirrhosis. Gut. 2010;59 (5 ):638–644. doi:10.1136/gut.2009.187286 19951909
11. Liu YI, Shin LK, Jeffrey RB, et al. Quantitatively defining washout in hepatocellular carcinoma. Am J Roentgenol. 2013;200 (1 ):84–89. doi:10.2214/AJR.11.7171 23255745
12. Zhou J, Zaiss M, Knutsson L, et al. Review and consensus recommendations on clinical APT-weighted imaging approaches at 3T: application to brain tumors. Magn Reson Med. 2022;88 (2 ):546–574. doi:10.1002/mrm.29241 35452155
• Outlines the current clinical APTw imaging approaches and provides a rationale for optimized APTw tumor imaging at 3 T. This consensus recommendations provides the first broadly accepted guidelines for APTw imaging of brain tumors on 3 T MRI systems from different vendors.

13. van Zijl PCM, Lam WW, Xu J, et al. Magnetization Transfer Contrast and Chemical Exchange Saturation Transfer MRI. Features and analysis of the field-dependent saturation spectrum. Neuroimage. 2018;168 :222–241. doi:10.1016/j.neuroimage.2017.04.045 28435103
14. Zhou J, Lal B, Wilson DA, et al. Amide proton transfer (APT) contrast for imaging of brain tumors. Magn Reson Med. 2003;50 (6 ):1120–1126. doi:10.1002/mrm.10651 14648559
• Seminal paper demonstrating the chemical exchange process governing APT imaging was more abundant in tumor tissue than in healthy normal tissue.

15. Zhou J, Zijl PCMv. Chemical exchange saturation transfer imaging and spectroscopy. Progress Nucl Magn Reson Spectrosc. 2006;48 (2–3 ):109–136. doi:10.1016/j.pnmrs.2006.01.001
• This reference highlights the sensitivity of CEST imaging/spectroscopy method describing signal enhancement factors of up to 106 for some systems.

16. Togao O, Kessinger CW, Huang G, et al. Characterization of lung cancer by amide proton transfer (APT) imaging: an in-vivo study in an orthotopic mouse model. PLOS ONE. 2013;8 (10 ):e77019. doi:10.1371/journal.pone.0077019 24143199
17. Sartoretti E, Sartoretti T, Wyss M, et al. Amide proton transfer weighted (APTw) imaging based radiomics allows for the differentiation of gliomas from metastases. Scient Rep. 2021;11 (1 ):5506. doi:10.1038/s41598-021-85168-8
18. Yang L, Wang L, Tan Y, et al. Amide Proton Transfer-weighted MRI combined with serum prostate-specific antigen levels for differentiating malignant prostate lesions from benign prostate lesions: a retrospective cohort study. Cancer Imag. 2023;23 (1 ):3. doi:10.1186/s40644-022-00515-w
19. Gao T, Zou C, Li Y, et al. A Brief History and Future Prospects of CEST MRI in Clinical Non-Brain Tumor Imaging. Inter J Mol Sci. 2021;22 (21 ):11559. doi:10.3390/ijms222111559
20. Sotirios B, Demetriou E, Topriceanu CC, et al. The role of APT imaging in gliomas grading: a systematic review and meta-analysis. Eur J Radiol. 2020;133 :109353. doi:10.1016/j.ejrad.2020.109353 33120241
21. Wu B, Jia F, Li X, et al. Comparative study of amide proton transfer imaging and intravoxel incoherent motion imaging for predicting histologic grade of hepatocellular carcinoma [original research]. Front Oncol. 2020;10 :562049. doi:10.3389/fonc.2020.562049 33194630
22. Zhou J, Tryggestad E, Wen Z, et al. Differentiation between glioma and radiation necrosis using molecular magnetic resonance imaging of endogenous proteins and peptides. Nat Med. 2011;17 (1 ):130–134. doi:10.1038/nm.2268 21170048
23. Nichelli L, Casagranda S. Current emerging MRI tools for radionecrosis and pseudoprogression diagnosis. Curr Opin Oncol. 2021;33 (6 ):597–607. doi:10.1097/CCO.0000000000000793 34534142
24. Ray KJ, Simard MA, Larkin JR, et al. Tumor pH and protein concentration contribute to the signal of amide proton transfer magnetic resonance imaging. Cancer Res. 2019;79 (7 ):1343–1352. doi:10.1158/0008-5472.CAN-18-2168 30679178
• Study elucidates the biological source of the APT MRI signal in tumors, thus advancing our understanding and interpretation of tumor APT MRI.

25. Reynolds L, Mulik RS, Wen X, et al. Low-density lipoprotein-mediated delivery of docosahexaenoic acid selectively kills murine liver cancer cells. Nanomedicine (Lond). 2014;9 (14 ):2123–2141. doi:10.2217/nnm.13.187 24397600
• Seminal paper which first describes formulation, characterization and the tumor selective cytotoxicity of LDL-DHA nanoparticles.

26. Gade TPF, Hunt SJ, Harrison N, et al. Segmental transarterial embolization in a translational rat model of hepatocellular carcinoma. J Vasc Interv Radiol. 2015;26 (8 ):1229–1237. doi:10.1016/j.jvir.2015.02.006 25863596
27. Ha WS, Kim CK, Song SH, et al. Study on mechanism of multistep hepatotumorigenesis in rat: development of hepatotumorigenesis. J Veter Sci. 2001;2 (1 ):53–58. doi:10.4142/jvs.2001.2.1.53
28. Ou W, Mulik RS, Anwar A, et al. Low-density lipoprotein docosahexaenoic acid nanoparticles induce ferroptotic cell death in hepatocellular carcinoma. Free Radic Biol Med. 2017;112 :597–607. doi:10.1016/j.freeradbiomed.2017.09.002 28893626
29. Heo HY, Zhang Y, Jiang S, et al. Quantitative assessment of amide proton transfer (APT) and nuclear overhauser enhancement (NOE) imaging with extrapolated semisolid magnetization transfer reference (EMR) signals: II. Comparison of three EMR models and application to human brain glioma at 3 Tesla. Magn Reson Med. 2016;75 (4 ):1630–1639. doi:10.1002/mrm.25795 26033553
30. Kurma K, Manches O, Chuffart F, et al. DEN-induced rat model reproduces key features of human hepatocellular carcinoma. Cancers. 2021;13 (19 ):4981. doi:10.3390/cancers13194981 34638465
• Describes how the rat model of DEN-induced hepatocellular carcinoma (HCC) mimics HCC seen in humans, including liver damage, chronic inflammation, hepatocytes proliferation, liver fibrosis, disorganized vasculature and altered liver immune microenvironment.

31. Lin Y, Luo X, Yu L, et al. Amide proton transfer-weighted MRI for predicting histological grade of hepatocellular carcinoma: comparison with diffusion-weighted imaging. Quant Imaging Med Surg. 2019;9 (10 ):1641–1651. doi:10.21037/qims.2019.08.07 31728308
32. Takayama Y, Nishie A, Togao O, et al. Amide proton transfer MR imaging of endometrioid endometrial adenocarcinoma: association with histologic grade. Radiology. 2018;286 (3 ):909–917. doi:10.1148/radiol.2017170349 29083987
33. Li G, Jiang G, Mei Y, et al. Applying amide proton transfer-weighted imaging (APTWI) to distinguish papillary thyroid carcinomas and predominantly solid adenomatous nodules: comparison with diffusion-weighted imaging [original research]. Front Oncol. 2020;10 :918. doi:10.3389/fonc.2020.00918 32637356
34. van Zijl PC, Zhou J, Mori N, et al. Mechanism of magnetization transfer during on-resonance water saturation. A new approach to detect mobile proteins, peptides, and lipids. Magn Reson Med. 2003;49 (3 ):440–449. doi:10.1002/mrm.10398 12594746
35. Dolfi SC, Chan LL-Y, Qiu J, et al. The metabolic demands of cancer cells are coupled to their size and protein synthesis rates. Cancer Metabol. 2013;1 (1 ):20. doi:10.1186/2049-3002-1-20
36. Lin J, Amir A. Homeostasis of protein and mRNA concentrations in growing cells. Nature Commun. 2018;9 (1 ):4496. doi:10.1038/s41467-018-06714-z 30374016
37. Xu J, Xie R, Liu X, et al. Expression and functional role of vacuolar H(+)-ATPase in human hepatocellular carcinoma. Carcinogenesis. 2012;33 (12 ):2432–2440. doi:10.1093/carcin/bgs277 22962303
38. Webb BA, Chimenti M, Jacobson MP, et al. Dysregulated pH: a perfect storm for cancer progression. Nat Rev Cancer. 2011;11 (9 ):671–677. doi:10.1038/nrc3110 21833026
39. Bhujwalla ZM, Aboagye EO, Gillies RJ, et al. Nm23-transfected MDA-MB-435 human breast carcinoma cells form tumors with altered phospholipid metabolism and pH: a 31P nuclear magnetic resonance study in vivo and in vitro. Magn Reson Med. 1999;41 (5 ):897–903. doi:10.1002/(SICI)1522-2594(199905)41:5<897::AID-MRM7>3.3.CO;2-K 10332871
40. Ding YF, Wu ZH, Wei YJ, et al. Hepatic inflammation-fibrosis-cancer axis in the rat hepatocellular carcinoma induced by diethylnitrosamine. J Cancer Res Clin Oncol. 2017;143 (5 ):821–834. doi:10.1007/s00432-017-2364-z 28238064
41. Connor F, Rayner TF, Aitken SJ, et al. Mutational landscape of a chemically-induced mouse model of liver cancer. J Hepatol. 2018;69 (4 ):840–850. doi:10.1016/j.jhep.2018.06.009 29958939
42. Hanna RF, Aguirre DA, Kased N, et al. Cirrhosis-associated hepatocellular nodules: correlation of histopathologic and MR imaging features. Radiographics. 2008;28 (3 ):747–769. doi:10.1148/rg.283055108 18480482
43. Sagiyama K, Mashimo T, Togao O, et al. In vivo chemical exchange saturation transfer imaging allows early detection of a therapeutic response in glioblastoma. Proc Natl Acad Sci U S A. 2014;111 (12 ):4542–4547. doi:10.1073/pnas.1323855111 24616497
44. Nishie A, Asayama Y, Ishigami K, et al. Amide proton transfer imaging to predict tumor response to neoadjuvant chemotherapy in locally advanced rectal cancer. J Gastroenterol Hepatol. 2019;34 (1 ):140–146. doi:10.1111/jgh.14315 29900583
45. Kumari N, Thakur N, Cho HR, et al. Assessment of early therapeutic response to nitroxoline in temozolomide-resistant glioblastoma by amide proton transfer imaging: a preliminary comparative study with diffusion-weighted imaging. Scientific Reports. 2019;9 (1 ):5585. doi:10.1038/s41598-019-42088-y 30944404
46. Wen X, Reynolds L, Mulik RS, et al. Hepatic arterial infusion of low-density lipoprotein docosahexaenoic acid nanoparticles selectively disrupts redox balance in hepatoma cells and reduces growth of orthotopic liver tumors in rats. Gastroenterology. 2016;150 (2 ):488–498. doi:10.1053/j.gastro.2015.10.008 26484708
